Adipose-derived stem cells and lattices

ABSTRACT

The present invention provides adipose-derived stem cells (ADSCs), adipose-derived stem cell-enriched fractions (ADSC-EF) and adipose-derivedlattices, alone and combined with the ADSCs of the invention. In one aspect, the present invention provides an ADSC substantially free of adipocytes and red blood cells and clonal populations of connective tissue stem cells. The ADSCs can be employed, alone or within biologically-compatible compositions, to generate differentiated tissues and structures, both in vivo and in vitro. Additionally, the ADSCs can be expanded and cultured to produce molecules such as hormones, and to provide conditioned culture media for supporting the growth and expansion of other cell populations. In another aspect, the present invention provides a adipose-derived lattice substantially devoid of cells, which includes extracellular matrix material from adipose tissue. The lattice can be used as a substrate to facilitate the growth and differentiation of cells, whether in vivo or in vitro, into anlagen or even mature tissues or structures.

[0001] This patent application is a continuation-in-part (CIP) of U.S.Ser. No. not yet known, filed Sep. 10, 2001, which corresponds to PCTapplication No. PCT/US00/06232, filed Mar. 10, 2000, which claims thebenfit of the filing dates of U.S. Ser. No. 60/123,711, filed Mar. 10,1999, and U.S. Ser. No. 60/162,462, filed Oct. 29, 1999. The contents ofall of the foregoing application are incorporated by refernce in theirentireties into the present patent application.

[0002] Throughout this application, various publications are referenced.The disclosures of these publications are hereby incorporated byreference herein in their entireties.

BACKGROUND OF THE INVENTION

[0003] In recent years, the identification of mesenchymal stem cells,chiefly obtained from bone marrow, has led to advances in tissueregrowth and differentiation. Such cells are pluripotent cells found inbone marrow and periosteum, and they are capable of differentiating intovarious mesenchymal or connective tissues. For example, such bone-marrowderived stem cells can be induced to develop into myocytes upon exposureto agents such as 5-azacytidine (Wakitani et al., Muscle Nerve, 18(12),1417-26 (1995)). It has been suggested that such cells are useful forrepair of tissues such as cartilage, fat, and bone (see, e.g., U.S. Pat.Nos. 5,908,784, 5,906,934, 5,827,740, 5,827,735), and that they alsohave applications through genetic modification (see, e.g., U.S. Pat. No.5,591,625). While the identification of such cells has led to advancesin tissue regrowth and differentiation, the use of such cells ishampered by several technical hurdles. One drawback to the use of suchcells is that they are very rare (representing as few as 1/2,000,000cells), making any process for obtaining and isolating them difficultand costly. Of course, bone marrow harvest is universally painful to thedonor. Moreover, such cells are difficult to culture without inducingdifferentiation, unless specifically screened sera lots are used, addingfurther cost and labor to the use of such stem cells. U.S. Pat. No.6,200,606 by Peterson et al., describes the isolation of CD34+ bone orcartilage precursor cells (of mesodermal origin) from tissues, includingadipose.

[0004] There remains a need for a more readily available source forlarge numbers of stem cells, particularly cells that can differentiateinto multiple lineages of different germ layers, and that can becultured without the requirement for costly prescreening of culturematerials.

[0005] Other advances in tissue engineering have shown that cells can begrown in specially-defined cultures to produce three-dimensionalstructures. Spacial definition typically is achieved by using variousacellular lattices or matrices to support and guide cell growth anddifferentiation. While this technique is still in its infancy,experiments in animal models have demonstrated that it is possible toemploy various acellular lattice materials to regenerate whole tissues(see, e.g., Probst et al. BJU Int., 85(3), 362-7 (2000)). A suitablelattice material is secreted extracellular matrix material isolated fromtumor cell lines (e.g., Engelbreth-Holm-Swarm tumor secretedmatrix—“matrigel”). This material contains type IV collagen and growthfactors, and provides an excellent substrate for cell growth (see, e.g.,Vukicevic et al., Exp. Cell Res, 202(1), 1-8 (1992)). However, as thismaterial also facilitates the malignant transformation of some cells(see, e.g., Fridman, et al., Int. J. Cancer, 51(5), 740-44 (1992)), itis not suitable for clinical application. While other artificiallattices have been developed, these can prove toxic either to cells orto patients when used in vivo. Accordingly, there remains a need for alattice material suitable for use as a substrate in culturing andgrowing populations of cells.

SUMMARY OF THE INVENTION

[0006] The present invention provides adipose-derived stem cells,adipose-derived stem cell fractions, lattices, and method for obtainingthe cells, fractions, and lattices. In one aspect, the present inventionprovides an adipose-derived stem cell fraction substantially free ofadipocytes and red blood cells and populations of connective tissuecells. The present invention also provides stem cells, isolated from thefraction, where the stem cells are pluripotent. The pluripotent stemcells have the ability to differentiate into mesoderm, ectoderm, orendoderm. The cells can be employed, alone or withinbiologically-compatible compositions, to generate differentiated tissuesand structures, both in vivo and in vitro. Additionally, the cells canbe expanded and cultured to produce growth factors and to provideconditioned culture media for supporting the growth and expansion ofother cell populations. In another aspect, the present inventionprovides a adipose-derived lattice substantially devoid of cells, whichincludes extracellular matrix material from adipose tissue. The latticecan be used as a substrate to facilitate the growth and differentiationof cells, whether in vivo or in-vitro, into anlagen or even maturetissues or structures.

[0007] Adipose tissue is plentiful and represent a ready source of thestem cells, fractions, and lattices. Moreover, the stem cells can bepassaged in culture in an undifferentiated state under cultureconditions not requiring prescreened lots of serum; the inventive cellscan be maintained with considerably less expense than other types ofstem cells. These and other advantages of the present invention, as wellas additional inventive features, will be apparent from the accompanyingdrawings and in the following detailed description.

BRIEF DESCRIPTION OF THE FIGURES

[0008]FIG. 1. Morphology; growth kinetics and senescence ofadipose-derived stem cells over long-term culture. Panel A: Themorphology of adipose-derived stem cells (e.g., a processed lipoaspirateor PLA) obtained from liposuctioned adipose tissue. Panel B:adipose-derived stem cells (PLAs) obtained from 3 donors, were culturedfor an extended period and cumulative population doubling was measuredand expressed as a function of passage number. Panel C: Senescence inadipose-derived stem cells (PLA) cultures as detected by staining forβ-galactosidase expression at pH 6.0. Representative senescent cells areshown (arrows).

[0009]FIG. 2. Composition of the adipose-derived stem cells (PLA) asdetermined by indirect immunofluorescence (IF). Adipose-derived stemcells (PLA) and bone marrow stromal cells (BMS), were stained with thefollowing antibodies: 1) anti-Factor VIII (FVIII); 2) anti-smooth muscleactin (SMA); and 3) ASO2 (ASO2). Factor VIII and smooth muscle actinexpressing cells are shown (arrows).

[0010]FIG. 3. Composition of the adipose-derived stem cells (PLA) asdetermined by flow cytometry. Panel A: Flow cytometry of adipose-derivedstem cells (PLA) samples using forward and side scatter (FS and SS,respectively). A representative adipose-derived stem cells sample isshown. Panel B: The cell composition of a representative adipose-derivedstem cells (PLA) sample from one donor was determined staining with thefollowing monoclonal antibodies: anti-Factor VIII (FVIII), anti-smoothmuscle actin (SMA), ASO2 and a monoclonal antibody to vimentin (VIM), anadditional marker for cells of mesenchymal origin. Panel C: Flowcytometry data from 5 donors was collected and the mean number ofpositive events for each cell-specific marker is expressed as apercentage of total adipose-derived stem cells (PLA) cell number.

[0011]FIG. 4. Adipose-derived stem cells (PLA) accumulate lipid-filleddroplets upon treatment with Adipogenic Medium (AM). Adipose-derivedstem cells (PLA), bone marrow-derived MSCs (MSC), and 3T3-L1pre-adipocyte cells (3T3-L1) were cultured for two weeks in AM andstained with Oil Red O to identify lipid-filled intracellular vacuoles.Undifferentiated PLA cells maintained in Control Medium (−ve Control)were stained as a negative control.

[0012]FIG. 5. Adipose-derived stem cells (PLA) induced with OsteogenicMedium (OM) express Alkaline Phosphatase and are associated with acalcified extracellular matrix (ECM). Adipose-derived stem cells (PLA),bone marrow-derived MSCs (MSC) and a human osteoblast cell line (NHOst)were cultured in OM to induce osteogenesis. Cells were stained at 2weeks for Alkaline Phosphatase activity (AP; red). The presence of acalcified extracellular matrix (black regions) was examined at 4 weeks(von Kossa). Undifferentiated adipose-derived stem cells maintained inControl Medium were examined for AP expression and matrix calcificationas a negative control (−ve Control).

[0013]FIG. 6. Adipose-derived stem cells (PLA) treated with ChondrogenicMedium (CM) are associated with a proteoglycan-rich matrix and expresscollagen type II. Adipose-derived stem cells (PLA) and MSCs (MSC) werecultured for 2 weeks in CM using the micromass technique to inducechondrogenesis. The cells were fixed and processed for the presence ofsulfated proteoglycans with Alcian Blue under acidic conditions (AlcianBlue). Paraffin sections of human cartilage were used as a positivecontrol (Cartilage) while undifferentiated PLAs maintained in ControlMedium were processed as a negative control (−ve Control). In addition,the expression of cartilage-specific collagen type II (Collagen II) wasexamined in PLA cells and human cartilage sections. Adipose-derived stemcells cultured in Control Medium (−ve Control) were stained with AlcianBlue and for collagen II expression as a negative control.

[0014]FIG. 7. Adipose-derived stem cells (PLA) cultured in MyogenicMedium (MM) express the myosin heavy chain and MyoD1. Adipose-derivedstem cells (PLA) were treated with MM and stained with antibodiesspecific to skeletal muscle myosin heavy chain (Myosin) or MyoD1(MyoD1). A human skeletal muscle cell line (SKM) was examined as apositive control. In addition, the presence of multinucleated cells inadipose-derived stem cells cultures is shown (PLA, inset box). Myosinand MyoD1 expression was also assessed in undifferentiatedadipose-derived stem cells (−ve Control) as a negative control.

[0015]FIG. 8. Growth kinetics of adipose-derived stem cells (PLA). PanelA: adipose-derived stem cells, isolated from each donor, were seeded intriplicate at a density of 1×10⁴ cells per well. Cell number wascalculated after 24 hours (day 1) and every 48 hours subsequent to day 1(days 3 through 11). Mean cell number for each donor was expressed withrespect to culture time. The growth curves from 4 representative donorsare shown (20 years—open squares, 39 years—open circles, 50 years—opentriangles and 58 years—crosses). Results are expressed as mean±SEM.Panel B: Population doubling was calculated in all donors from the logphase of each growth curve (i.e. from day 3 to day 9) and expressedaccording to age. The line of regression was calculated (n=20; r=0.62)

[0016]FIG. 9. Histological confirmation of adipogenic and osteogenicdifferentiation by adipose-derived stem cells (PLA). A: To confirmadipogenesis, cells were stained at 2 weeks post-induction with Oil RedO. Low and extensive adipogenic levels are shown (Panel 1—low; Panel2—high). Adipose-derived stem cells cultured in non-inductive controlmedium were analyzed as negative controls (Panel 3). B: To quantifyadipogenic differentiation, the number of Oil Red O-positive stainedcells were counted within three defined regions. Two samples wereanalyzed from each donor. The mean number of Oil Red O-positive cellswas determined and expressed as a percentage of total adipose-derivedstem cells number as an indication of adipogenic differentiation.Differentiation was expressed with respect to age and the line ofregression calculated (n=20; r=0.016).

[0017]FIG. 10. Osteogenic differentiation decreases with increasingdonor age. Panel A: To confirm osteogenesis, adipose-derived stem cells(PLA) were stained at 2 weeks post-induction for alkaline phosphotase(AP) activity (Panels 1 to 3) and at 4 weeks post-induction for matrixcalcification using von Kossa staining (Panels 4 to 6). Osteogenicdifferentiation levels are shown (Panels 1/2—low; Panels 4/5—high).Adipose-derived stem cells cultured in non-inductive control medium wereanalyzed as negative controls (Panels 3 and 6). Panel B: To quantifyosteogenic differentiation, the number of AP-positive stained cells werecounted within three defined regions. Two samples were analyzed fromeach donor. The mean number of AP-positive cells was determined andexpressed as a percentage of total adipose-derived stem cells number asan indication of the osteogenic differentiation. Differentiation wasexpressed with respect to age and the line of regression calculated(n=18; r=−0.70). Panel C: Based on the results of Panel B, the donorpool was divided into two age groups [(20 to 36 years (n=7) and 37 to 58years (n=11)]. The average level of osteogenic differentiation wascalculated for each group and expressed as a percentage of totaladipose-derived stem cells number. Statistical significance wasdetermined using an unpaired student t test assuming unequal variances(p<0.00 1). Differentiation is expressed as mean±SEM.

[0018]FIG. 11. Osteoprogenitor cell number within an adipose-derivedstem cell fraction (PLA fraction) does not significantly change withage. Osteoprogenitor cell number within the fraction was determined byidentifying cells with osteogenic potential. Two groups of donors wereexamined [Group A=20 to 39 years (n=5), Group B=40-58 years (n=6)].Osteogenesis was confirmed by staining for AP activity. Coloniescontaining more than 10 AP-positive cells (CFU/AP⁺) were counted andaveraged as an indicator of the number of osteogenic precursors withineach age group. Statistical significance was determined using anunpaired student t test assuming unequal variances (p=0.11). Values areexpressed as mean CFU/AP⁺±SEM.

[0019]FIG. 12. Human adipose-derived stem cells (PLA) placed inmicromass cultures and induced with chondrogenic media undergo cellularcondensation and nodule formation. Adipose-derived stem cells inducedunder micromass conditions were stained with Alcian blue staining at pH1 to detect the presence of sulfated proteoglycans. Panel A: cellularcondensation; (Panel B) ridge formation; (Panel C) formation ofthree-dimensional spheroids are shown (magnification 100×); (Panel D)negative control (control medium).

[0020]FIG. 13. Hematoxylin & Eosin, Goldner's trichrome, and Alcian bluestaining of nodule paraffin sections from adipose-derived stem cells(PLA). Micromass cultures adipose-derived stem cells were treated withchondrogenic medium to form nodules, the nodules were embedded inparaffin and sectioned. Nodule sections were stained using conventionalhematoxylin and eosin (Panels A and B) and a Goldner's trichrome stainto detect collagens (green) (Panels C and D). Adipose-derived stem cellsinduced for 2 days are shown at a magnification of 200× (Panels A and C)and 14 days are shown at 100× (Panels B and D). In addition, sectionswere stained with Alcian blue staining at pH 1, to detect highlysulfated proteoglycans. Day two nodules (Panel E) are shown at amagnification of 200× and day fourteen nodules (Panel F) are shown at100×.

[0021]FIG. 14. Nodule differentiated from adipose-derived stem cells(PLA) express chondroitin-4-sulfate and keratin sulfate as well ascartilage-specific collagen type II. Nodules induced fromadipose-derived stem cells for 2 days (Panels A and C) and 14 days(Panels B and D) were embedded in paraffin and sectioned. Sections werestained with monoclonal antibodies to the sulfated proteoglycanschondroitin-4-sulfate and keratin sulfate. Sections were also stainedwith monoclonal antibodies to collagen type II (Panels E and F)(magnification 200×).

[0022]FIG. 15. RT-PCR analysis of nodules induced from adipose-derivedstem cells confirms the expression of collagens type II and type X aswell as expression of cartilage-specific proteoglycan and aggrecan.Adipose-derived stem cells induced for 2, 7, and 14 days in chondrogenicmedium and non-inductive control medium were analyzed by RT-PCR for theexpression of collagen type I (CN I), type II (CN II), and type X (CN X)as well as cartilage-specific proteoglycan (PG), aggrecan (AG), andosteocalcin (OC).

[0023]FIG. 16. Adipose-derived stem cells induced in Myogenic Mediumexpress MyoD1. Panels A to C: adipose-derived stem cells (PLA) werestained with an antibody to MyoD1 following 1 week (Panel A), 3 weeks(Panel B) and 6 weeks (Panel C) induction in MM. Expression of MyoD1 inthe nucleus of positive staining PLA cells is shown (arrows,magnification 200×). Panels D to F: PLA cells induced for 1 week (PanelD), 3 weeks (Panel E) and 6 weeks (Panel F) in non-inductive controlmedium (CM) were processed as above as a negative control (magnification200×).

[0024]FIG. 17. Adipose-derived stem cells induced in Myogenic Mediumexpress skeletal muscle myosin heavy chain. Panels A to C:adipose-derived stem cells (PLA) cells were stained with an antibody tothe myosin heavy chain (myosin) following 1 week (Panel A), 3 weeks(Panel B) and 6 weeks (Panel C) induction in MM. Myosin-positivestaining PLA cells are shown (arrows, magnification 200×). Panels D toF: adipose-derived stem cells (PLA) cells induced for 1 week (Panel D),3 weeks (Panel E) and 6 weeks (Panel F) in non-inductive CM wereprocessed as above as a negative control (magnification 200×).

[0025]FIG. 18. Adipose-derived stem cells cultured in Myogenic Mediumform multi-nucleated cells. Panel A: Phase contrast of adipose-derivedstem cells (PLA) at 3 weeks (1) and 6 weeks (2) post-induction with MM(magnification 400×). Multi-nucleated cells are shown (arrows). Panel B:Immunostaining of adipose-derived stem cells (PLA) cells at 6 weekspost-induction with an antibody to the myosin heavy chain.Myosin-expressing multi-nucleated cells are shown (arrows).

[0026]FIG. 19: RT-PCR analysis of adipose-derived stem cells induced inMM. RT-PCR was performed on adipose-derived stem cells induced for 1, 3and 6 weeks in MM (PLA-MM) or in CM (PLA-CM), using primers to humanMyoD1 and myosin. RT-PCR analysis of human foreskin fibroblast (HFF)cells induced in MM (HFF-MM) was also performed as a negative control.Duplicate reactions were performed using a primer set to β-actin as aninternal control. PCR products were resolved by agarose gelelectrophoresis and equalized using β-actin levels.

[0027]FIG. 20. The proportion of MyoD1-positive adipose-derived stemcells increases with induction time. Histogram showing the mean numberof MyoD1-positive, adipose-derived stem cells (PLA) after a 1, 3 and 6week induction in MM (% of total PLA cells±SEM—hatched bars). The meannumber of MyoD1-positive cells observed after induction ofadipose-derived stem cells with CM (black bars) and HFF cells in MM(open bars) was also measured. The values for each experiment are shownin table format below. A statistical comparison of MyoD1 values from 1to 6 weeks using a one-way ANOVA was performed (asterisks; P<0.001,F=18.9). Furthermore, an ANOVA was performed comparing the experimentaland control values for each time point. The p-values are shown(p<0.0001).

[0028]FIG. 21. A time-dependent increase in myosin expression isobserved in induced adipose-derived stem cells. Histogram showing themean number of myosin-positive adipose-derived stem cells (PLA) after a1, 3 and 6 week induction in myosin medium (MM) (% of total PLAcells±SEM—hatched bars). The mean number of myosin-positive cellsobserved after induction of adipose-derived stem cells with controlmedium (CM) (black bars), and human foreskin fibroblast cells (HFF) inmyosin medium (MM) (open bars) was also measured. The values for eachexperiment are shown in table format below. A statistical comparison ofmyosin values from 1 to 6 weeks using a one-way ANOVA was performed(asterisks; P<0.0001, F=75.5). Furthermore, an ANOVA was performedcomparing the experimental and control values for each time point. Thep-values are shown (p<0.0001).

[0029]FIG. 22. Long-term chrondrogenic potetial of adipose-derived stemcells. Adipose-derived stem cells, at passage 1 (panel A), 3 (panel B),and 15 (panel C), were induced under micromass conditions and stainedwith Alcian blue staining at pH 1 to detect the presence of sulfatedproteoglycans.

[0030]FIG. 23. The adipose-derived stem cells (PLA) express a unique setof CD markers. PLA cell and MSCs from human bone marrow were processedfor IF for the indicated CD antigens. Cells were co-stained with DAPI tovisualize nuclei (blue) and the fluorescent images combined.

[0031]FIG. 24. CD marker profile of adipose-derived stem cells (PLA) andbone marrow MSCs using flow cytometry. Panel A: Adipose-derived stemcells were analyzed by FC using forward and side scatter to assess cellsize and granularity (FSC-H and SSC-H, respectively). MSCs were analyzedas a control. Panel B: PLA cells were fixed and incubated for theindicated CD markers using fluorochrome-conjugated primary antibodies.Stained PLA cells were subsequently analyzed by FC. MSCs and PLA cellsstained with fluorochrome-conjugated non-specific IgG were examined as apositive and negative control, respectively. All results were correctedfor senescence and represent a total of 10⁵ events.

[0032]FIG. 25. Osteogenic adipose-derived stem cells (PLA) can becharacterized by distinct proliferative, synthetic and mineralizationphases. Adipose-derived stem cells were harvested and plated into 35 mmtissue culture dishes in two sets of four plates per differentiationperiod. All dishes were maintained in Control medium until approximately50% confluence was reached. The cells were induced with Osteogenicmedium (OM) and cell number was counted at the indicated days. Cellnumber was expressed as the number of adipose-derived stem cells (#cells (10⁵ )) and plotted versus differentiation time (Panel A). Foreach time period, one dish was stained for alkaline phosphatase (AP)activity and one dish was stained using a Von Kossa stain (VK) to detectcalcium phosphate (Panel B).

[0033]FIG. 26. Dexamethasone and 1,25-dihydroxyvitamin D₃ differentiallyaffect PLA osteogenesis: AP enzyme and calcium phosphate quantitation.Triplicate samples of PLA cells, MSCs and NHOsts were induced for up to6 weeks in OM, containing either 10⁻⁷ M Dexamethasone (OM/Dex) or 10⁻⁸ M1,25-dihydroxyvitamin D₃ (OM/VD). Cells were assayed for AP activity,total calcium content and total protein. AP levels were expressed asnmol p-nitrophenol formed per minute per microgram protein (nmolp-nitrophenol/min/ug). Calcium levels were expressed as mM calcium permicrogram protein (mM Ca²⁺/ug). Non-induced PLA cells (Control) wereanalyzed as a negative control. Values were expressed as the mean±SD.

[0034]FIG. 27. Osteo-induced PLA cells express several genes consistentwith osteogenic differentiation: RT-PCR and Microarray analyses. PanelA: PLA cells were cultured in either OM/Dex, OM/VD or non-inductiveControl medium (Control) for the indicated days. Total RNA was isolated,cDNA synthesized and PCR amplification performed for the indicatedgenes. MSCs were induced in OM/Dex or OM/VD and NHOsts were induced for2 and 3 weeks in OM/Dex as controls. Duplicate reactions were amplifiedusing primers to β-actin as an internal control. Panel B: PLA cells wereinduced for 3 weeks in OM/Dex or maintained in non-inductive controlmedium. Total RNA was isolated and subject to microarray analysis usinga customized array containing the genes, OC, OP, ON, CBFA1, CNI and BSP.

[0035]FIG. 28. Osteo-induced PLA cells express several proteinsconsistent with osteogenic differentiation: Immunofluorescent andWestern analyses. Panel A: PLA cells and MSCs were induced in OM/Dex ormaintained in non-inductive Control medium (Control) for 21 days. Cellswere processed for IF for the expression of OC, OP and ON. Cells wereco-stained with DAPI to visualize nuclei (blue) and the fluorescentimages combined. Panel B: PLA cells were cultured in OM/Dex ornon-inductive Control medium (Control) for 7 and 21 days. Cell lysateswere separated by electrophoresis and analyzed by Western blotting usingantibodies to OP (αOP), ON (αON), Decorin (αDEC), Biglycan (αBG) and CNI(αCNI). The expression of the transferrin receptor (αTfR) was used as aninternal control.

[0036]FIG. 29. Adipogenic differentiation by adipose-derived stem cells(PLA) is accompanied by growth arrest. Adipose-derived stem cells wereharvested and plated into 35 mm tissue culture dishes in one set of fourplates per differentiation period. All dishes were maintained in Controlmedium until approximately 80% confluence was reached. The cells wereinduced with Adipogenic medium (AM) and cell number was counted at theindicated days. Cell number was expressed as the number of PLA cells (#cells (10⁵)) and plotted versus differentiation time (Panel A). For eachtime period, one dish was stained with Oil Red O to detect lipidaccumulation (Panel B).

[0037]FIG. 30. Adipogenic PLA cells express GPDH activity. Triplicatesamples of PLA cells and 3T3-L1 cells were induced for up to 5 weeks inAM (PLA—AM, 3T3—AM, respectively). The cells were assayed for GPDHactivity and total protein. GPDH levels were expressed as units GPDH permicrogram protein (GPDH/ug). Non-induced PLA cells were analyzed as anegative control (PLA—Control). Values were expressed as mean±SD.

[0038]FIG. 31. Adipose-derived stem cells express several genesconsistent with adipogenic differentiation: RT-PCR: Adipose-derived stemcells were induced in AM (AM) or maintained in non-inductive Controlmedium (Control) for the indicated days. Cells were analyzed by RT-PCRfor the indicated genes. MSCs and 3T3-L1 cells were induced in AM ascontrols. Duplicate reactions were amplified using primers to β-actin asan internal control.

[0039]FIG. 32. Adipose-derived stem cell induced toward the chondrogeniclineage are associated with the proteoglycans keratan and chondroitinsulfate: Immunohistochemistry and Dimethyldimethylene blue assay. PanelA: Adipose-derived stem cells (PLA), under micromass conditions, wereinduced in chondrogenic medium (CM) or maintained in non-inductiveControl medium (Control) for 7 days. Nodules were fixed, embedded inparaffin, sectioned and stained with Alcian Blue to identify sulfatedproteoglycans. Sections were also stained for the expression of CNII,keratan sulfate (KS) and chondroitin-4-sulfate (CS), followed bycounter-staining using H&E. Panel B: Triplicate samples of PLA cells andNHCK cells were induced for up to 3 weeks in CM (PLA—CM, NHCK—CM,respectively). Proteoglycan levels (keratan sulfate and chondroitinsulfate) were determined and expressed as microgram proteoglycan permicrogram total protein (ug PG/ug). Non-induced, Adipose-derived stemcells (PLA—Control) were analyzed as a negative control. Values wereexpressed as the mean±SD.

[0040]FIG. 33. Chondrogenic PLA cells express several genes consistentwith cartilage differentiation: RT-PCR. PLA cells, under micromassculture conditions, were induced in CM for 4, 7, 10 and 14 days ormaintained in non-inductive Control medium for 10 days (Control). Cellswere analyzed by RT-PCR for the indicated genes. NHCK cells were inducedin a commercial pro-chondrogenic medium as a positive control. Duplicatereactions were performed using primers to β-actin as an internalcontrol.

[0041]FIG. 34. PLA cells induced toward the myogenic lineage expressseveral genes consistent with myogenic differentiation: RT-PCR analysis.PLA cells were induced in MM (PLA—MM) for 1, 3 and 6 weeks. Cells wereanalyzed by RT-PCR for the expression of MyoD1 (MD1), myosin (MYS),myogenin (MG) and myf5 (MYF5). Total RNA prepared from human skeletalmuscle (SKM) was analyzed as a positive control. Duplicate reactionswere amplified using primers to β-actin as an internal control.

[0042]FIG. 35. ADSCs express multiple markers consistent withmulti-lineage capacity. ADSC Isolation: PLA cells were plated atextremely low confluency in order to result in isolated single cells.Cultures were maintained in Control medium until proliferation of singlePLA cells resulted in the formation of well-defined colonies. The singlePLA-cell derived colonies were termed Adipose Derived Stem Cells(ADSCs). ADSCs were harvested using sterile cloning rings and 0.25%trypsin/EDTA. The harvested ADSCs were amplified in Cloning Medium (15%FBS, 1% antibiotic/antimycotic in F12/DMEM (1:1)). Tri-lineage ADSCclones were differentiated in OM, AM and CM and multi-lineage capacityby IH using the following histological and IH assays: AlkalinePhosphatase (osteogenesis), Oil Red O (adipogenic) and Alcian Blue(chondrogenic).

[0043]FIG. 36. Isolation of multi-lineage clones from PLA populationsdoes not alter the expression profile of CD markers. Dual- andtri-lineage clones were isolated and expanded from single PLA cells. Theclone populations were processed for the expression of the indicated CDmarkers using IF. The ADSCs were co-stained with DAPI to visualizenuclei (blue) and the fluorescent images combined.

[0044]FIG. 37. ADSCs express multiple genes consistent withmulti-lineage capacity. Tri-lineage ADSC clones were cultured in OM/VD(ADSC—Bone), AM (ADSC—Fat) and CM (ADSC—Cartilage), in addition tocontrol medium (ADSC—Control), followed by RT-PCR analysis for theindicated lineage-specific genes. β-actin levels were analyzed as aninternal control.

[0045]FIG. 38. PLA cells appear to exhibit neurogenic capacity in vitro.Panel A: Light micrographs of non-induced PLA cells (PLA—0 hrs) and PLAcells induced with NM for 2 and 8 hrs (PLA—2hrs, PLA—8 hrs,respectively). Panel B: PLA cells were maintained in NM or Controlmedium for 5 hours (PLA—NM, PLA—Control, respectively) and analyzed byIH for expression of the following lineage-specific markers: NSE, trk-A,NeuN and MAP-2 (neural), GFAP (astrocytic). PC12 cells treated with NGFwere also assessed as a positive control. Panel C: PLA cells wereinduced in NM for 4.5 and 9 hrs and analyzed by RT-PCR for the indicatedgenes. In addition, PLA cells were induced in NM for 9 hrs andmaintained in NPMM for 1 week (NPMM). Non-induced PLA cells (Control)were analyzed as a negative control. PC12 cells were examined as apositive control, together with total RNA prepared from human brain(Brain).

[0046]FIG. 39. Clones isolated from adipose-derived stem cell fractionsexhibit neurogenic potential. Clones were examined usingimunnohistochemistry for adipogenic (oil red O stain), osteogenic(alkaline phosphotase), chondrogenic (Alcian blue stain), and neurogenic(anti-trka expression) differntiation.

[0047]FIG. 40. Osteogenic differentiation of the adipose-derived stemcells (PLA) does not significantly alter CD marker expression. PLA cells(Panel A) and MSCs (Panel B) were induced in OM for 3 weeks (PLA—Bone,MSC—Bone respectively), or maintained in non-inductive Control medium(PLA—Control, MSC—Control). Cells were processed for IF for theexpression of CD34, CD44, CD45 and CD90, co-stained with DAPI tovisualize nuclei (blue) and the fluorescent images combined.

[0048]FIG. 41. Adipogenic differentiation results in subtle changes tothe adipose-derived stem cells (PLA) CD marker profile. PLA cells (PanelA) and MSCs (Panel B) were induced in AM for 2 weeks (PLA—Fat, MSC—Fat,respectively) or maintained in non-inductive Control medium(PLA—Control, MSC—Control). Cells were processed for IF for theexpression of CD34, CD44, CD45 and CD90, co-stained with DAPI tovisualize nuclei (blue) and the fluorescent images combined. Tovisualize adipocytes and their staining pattern, fluorescent images werecombined with light micrographs (inset). Lipid-filled cells (whitearrows—fluorescent image; black arrows—inset) and fibroblasts (filledwhite arrows—fluorescent image; filled black arrows—inset) areindicated.

[0049]FIG. 42. Differentiation alters the expression of specific CDmarkers on adipose-derived stem cells (PLA): Flow cytometry. Panel A:PLA cells were maintained for 2 weeks in Control medium (Control), or inOM (Osteogenic) or AM (Adipogenic). Cells were analyzed by FC usingforward and side scatter to assess cell size and granularity (FSC-H andSSC-H, respectively). Panels B and C: PLA cells were maintained for 2weeks in Control medium (PLA—CM), or in OM (PLA—OM) or AM (PLA—AM).Cells were directly stained for the indicated CD markers usingfluorochrome-conjugated primary antibodies and analyzed by FC. Theadipose-derived stem cells, stained with fluorochrome-conjugatednon-specific IgG, were examined as a negative control. All results werecorrected for senescence and represent a total of 10⁵ events.

[0050]FIG. 43. Differentiation of the adipose-derived stem cells (PLA)results in a change in ECM composition. PLA cells were induced foreither 3 weeks in OM (PLA—Bone), 2 weeks in AM (PLA—Fat) or maintainedin Control medium (PLA—Control). Cells were processed for IF usingantibodies to collagen type 1 (CNI), type 4 (CNIV) and type 5 (CNV).Cells were co-stained with DAPI to visualize nuclei (blue) and thefluorescent images combined. Fluorescent images were combined with lightmicrographs (inset). Lipid-filled PLA cells (white arrows—fluorescentimage; black arrows—inset) are indicated. Osteo-induced MSCs (MSC—Bone),adipo-induced MSCs (MSC—Fat) and non-induced MSCs (MSC—Control) werealso analyzed.

DETAILED DESCRIPTION OF THE INVENTION Definitions

[0051] As used herein, “stem cell” defines an adult undifferentiatedcell that can produce itself and a further differentiated progeny cell.

[0052] As used herein, the “lineage” of a cell defines the heredity ofthe cell, i.e.; which cells it came from and what cells it can give riseto. The lineage of a cell places the cell within a hereditary scheme ofdevelopment and differentiation.

[0053] As used herein, the term “differentiates or differentiated”defines a cell that takes on a more committed (“differentiated”)position within the lineage of a cell. “Dedifferentiated” defines a cellthat reverts to a less committed position within the lineage of a cell.

[0054] As used herein, “a cell that differentiates into a mesodermal (orectodermal or endodermal) lineage” defines a cell that becomes committedto a specific mesodermal, ectodermal or endodermal lineage,respectively. Examples of cells that differentiate into a mesodermallineage or give rise to specific mesodermal cells include, but are notlimited to, cells that are adipogenic, chondrogenic, cardiogenic,dermatogenic, hematopoetic, hemangiogenic, myogenic, nephrogenic,urogenitogenic, osteogenic, pericardiogenic, or stromal.

[0055] Examples of cells that differentiate into ectodermal lineageinclude, but are not limited to epidermal cells, neurogenic cells, andneurogliagenic cells.

[0056] Examples of cells that differentiate into endodermal lineageinclude, but are not limited to pleurigenic cells, and hepatogeniccells, cell that give rise to the lining of the intestine, and cellsthat give rise to pancreogenic and splanchogenic cells.

[0057] As used herein, a “pluripotent cell” defines a lessdifferentiated cell that can give rise to at least two distinct(genotypically and/or phenotypically) further differentiated progenycells.

[0058] A “multi-lineage stem cell” or “multipotent stem cell” refers toa stem cell that reproduces itself and at least two furtherdifferentiated progeny cells from distinct developmental lineages. Thelineages can be from the same germ layer (i.e. mesoderm, ectoderm orendoderm), or from different germ layers. An example of two progenycells with distinct developmental lineages from differentiation of amulti-lineage stem cell is a myogenic cell and an adipogenic cell (bothare of mesodermal origin, yet give rise to different tissues). Anotherexample is a neurogenic cell (of ectodermal origin) and adipogenic cell(of mesodermal origin).

[0059] As used here, “adipose tissue” defines a diffuse organ of primarymetabolic importance made-up of white fat, yellow fat or brown fat. Theadipose tissue has adipocytes and stroma. Adipose tissue is foundthroughout the body of an animal. For example, in mammals, adiposetissue is present in the omentum, bone marrow, subcutaneous space andsurrounding most organs.

[0060] As used herein “conditioned media” defines a medium in which aspecific cell or population of cells have been cultured in, and thenremoved. While the cells were cultured in said medium, they secretecellular factors that include, but are not limited to hormones,cytokines, extracellular matrix (ECM), proteins, vesicles, antibodies,and granules. The medium plus the cellular factors is the conditionedmedium.

[0061] As used herein “isolated” defines a substance, for example anadipose-derived stem cell, that is separated from contaminants (i.e.substances that differ from the stem cell).

[0062] The present invention provides adipose-derived stem cells (ADSCs)and methods for obtaining them from a mesodermal origin (e.g., adiposetissue) and using them. Surprisingly, the inventive ADSCs candifferentiate into cells that give rise to more than one type of germlayer, e.g. mesoderm, endoderm or ectoderm, and combinations thereof,and are thus “multilineage” or “multipotent” cells.

[0063] In another embodiment, the ADSCs can differentiate into two ormore distinct lineages from different germ layers (such as endodermaland mesodermal), for example hepatocytes and adipocytes.

[0064] The ADSCs of the invention can differentiate into cells of two ormore lineages, for example adipogenic, chondrogenic, cardiogenic,dermatogenic, hematopoetic, hemangiogenic, myogenic, nephrogenic,neurogenic, neuralgiagenic, urogenitogenic, osteogenic, pericardiogenic,peritoneogenic, pleurogenic, splanchogenic, and stromal developmentalphenotypes. While such cells can retain two or more of these differentlinages (or developmental phenotypes), preferably, such ADSCs candifferentiate into three or more different lineages. The most preferredADSCs can differentiate into four or more lineages.

[0065] The ADSCs of the invention have the capacity to differentiateinto mesodermal tissues, such as mature adipose tissue, bone, varioustissues of the heart (e.g., pericardium, epicardium, epimyocardium,myocardium, pericardium, valve tissue, etc.), dermal connective tissue,hemangial tissues (e.g., corpuscles, endocardium, vascular epithelium,etc.), hematopeotic tissue, muscle tissues (including skeletal muscles,cardiac muscles, smooth muscles, etc.), urogenital tissues (e.g.,kidney, pronephros, meta- and meso-nephric ducts, metanephricdiverticulum, ureters, renal pelvis, collecting tubules, epithelium ofthe female reproductive structures (particularly the oviducts, uterus,and vagina), mesodermal glandular tissues (e.g., adrenal cortextissues), and stromal tissues (e.g., bone marrow). Of course, inasmuchas the ADSC can retain potential to develop into a mature cell, it alsocan realize its developmental phenotypic potential by differentiatinginto an appropriate precursor cell (e.g., a preadipocyte, a premyocyte,a preosteocyte, etc.).

[0066] In another embodiment, the ADSCs have the capacity todifferentiate into ectodermal tissues, such as neurogenic tissue, andneurogliagenic tissue.

[0067] In another embodiment, the ADSCs have the capacity todifferentiate into endodermal tissues, such as pleurogenic tissue, andsplanchnogenic tissue, and hepatogenic tissue, and pancreogenic tissue.

[0068] In yet another embodiment, ADSCs have the capacity todedifferentiate into developmentally immature cell types. Examples ofADSCs dedifferentiating into an immature cell type, include embryoniccells and fetal cells.

[0069] In another embodiment, the inventive ADSCs can give rise to oneor more cell lineages from one or more germ layers such as neurogeniccells (of ectodermal origin) and myogenic cells (of mesodermal origin).

[0070] The inventive ADSCs are useful for tissue engineering, woundrepair, in vivo and ex vivo tissue regeneration, tissue transplantation,and other methods that require cells that can differentiate into avariety of phenotypes and genotypes, or can support other cell types invivo or in vitro.

[0071] One aspect of the invention pertains to an adipose-derived stemcell-enriched fraction (ADSC-EF) that contains adipose-derived stemcells (ADSCs) of the invention. Preferably, the ADSC-EF is substantiallyfree of other cell types (e.g., adipocytes, red blood cells, and otherstromal cells, etc.) and extracellular matrix material. More preferably,the ADSC-EF is completely free of such other cell types and matrixmaterial. The ADSC-EF is obtained from adipose tissue of a mammal. Thepreferred embodiment includes an ADSC-EF obtained from adipose tissue ofa higher primate (e.g., a baboon or ape). The most preferred ADSCenriched fraction is obtained from human adipose tissue, using themethods described herein.

Methods of Obtaining ADSC-EF and ADSCs of the Invention

[0072] The ADSCs of the invention are isolated from adipose tissue. Theadipose tissue can be obtained from an animal by any suitable method. Afirst step in any such method requires the isolation of the adiposetissue from the source animal. The animal can be alive or dead, so longas adipose stromal cells within the animal are viable. Typically, humanadipose tissue is obtained from a living donor, using well-recognizedprotocols such as surgical or suction lipectomy. The preferred method toobtain human adipose tissue is by excision or liposuction procedureswell known in the art. Preferably, the inventive ADSCs are isolated froma liposuction aspirate. The ADSCs of the invention are present in theinitially excised or extracted adipose tissue, regardless of the methodby which the adipose tissue is obtained.

[0073] Three deposits of lipoaspirates, each from a different patient,identified as 1′,2′,3′, have been deposited on Sep. 7, 2001, with theAmerican Type Culture Collection (ATCC), 10801 University Blvd.,Manassas, Va. 20110-2209, under the provisions of the Budapest Treaty,and have been accorded ATCC deposit numbers ______, ______ and ______.

[0074] However obtained, the adipose tissue is processed to separate theADSCs of the invention from the remainder of the adipose tissue. TheADSC-EF that contains the ADSCs is obtained by washing the obtainedadipose tissue with a physiologically-compatible solution, such asphosphate buffer saline (PBS). The washing step consists of rinsing theadipose tissue with PBS, agitating the tissue, and allowing the tissueto settle. In addition to washing, the adipose tissue is dissociated.The dissociation can occur by enzyme degradation and neutralization.Alternatively, or in conjunction with such enzymatic treatment, otherdissociation methods can be used such as mechanical agitation, sonicenergy, or thermal energy. Three layers form after the washing,dissociation, and settling steps. The top layer is a free lipid layer.The middle layer includes the lattice and adipocyte aggregates. Themiddle layer is referred to as an “adipose-derived lattice enrichedfraction.” The middle layer or the lattice-enriched fraction is filteredto concentrate the lattice of the invention. A method of filtrationinvolves passing the middle layer through a large pore filter. Thematerial which does not pass through the filter includes the inventivelattice and aggregates of adipocytes. The adipose-derived lattice can bemanually separated from the other cells which did not pass through thefilter.

[0075] The bottom layer contains the ADSC-EF and the inventive ADSCs.The bottom layer is further processed to isolate the ADSCs of theinvention. The cellular fraction of the bottom layer is concentratedinto a pellet. One method to concentrate the cells includescentrifugation.

[0076] The bottom layer is centrifuged and the pellet is retained. Thepellet is designated the adipose-derived stem cell-enriched fraction(ADSC-EF) which includes the adipose-derived stem cell-enriched fraction(ADSC-EF). The ADSC-EF may contain erythrocytes (RBCs). In a preferredmethod the RBCs are lysed and removed. Methods for lysis and removedRBCs are well known in the art (e.g., incubation in hypotonic medium).If the RBCs are removed, then the RBC-free fraction contains the ADSC-EFfraction and the ADSCs. However, the RBCs are not required to be removedfrom the ADSC-EF.

[0077] The pellet is resuspended and can be washed (in PBS),centrifuged, and resuspended one or more successive times to achievegreater purity of the ADSCs. The ADSC-EF of the invention maybe aheterogenous population of cells which include the ADSCs of theinvention and adipocytes. The cells of the washed and resuspended pelletare ready for plating.

[0078] The ADSCs in the resuspended pellet can be separated from othercells of the resuspended pellet by methods that include, but are notlimited to, cell sorting, size fractionation, granularity, density,molecularly, morphologically, and immunohistologically.

[0079] In one embodiment, the ADSCs are separated from the other cellson the basis of cell size and granularity where ADSCs are small andagranular. Alternatively, a molecular method for separating the ADSCsfrom the other cells of the pellet is by assaying the length of thetelomere. Stem cells tend to have longer telomeres than differentiatedcells.

[0080] In another embodiment, a biochemical method for separating theADSCs from the other cells of the pellet is used by assaying telomeraseactivity. Telomerase activity can serve as a stem cell-specific marker.

[0081] In still another embodiment, the ADSCs are separated from theother cells of the pellet immunohistochemically, for example, bypanning, using magnetic beads, or affinity chromatography.

[0082] Alternatively, the process of isolating the ADSC enrichedfraction with the ADSCs is with a suitable device, many of which areknown in the art (see, e.g., U.S. Pat. No. 5,786,207). Such devices canmechanically achieve the washing and dissociation steps.

Culturing ADSCs

[0083] The ADSCs in the ADSC-EF can be cultured and, if desired, assayedfor number and viability, to assess the yield.

[0084] In one embodiment, the stem cells are cultured withoutdifferentiation using standard cell culture media (e.g., DMEM, typicallysupplemented with 5-15% (e.g., 10%) serum (e.g., fetal bovine serum,horse serum, etc.). Preferably, the stem cells are passaged at leastfive times in such medium without differentiating, while still retainingtheir developmental phenotype, and more preferably, the stem cells arepassaged at least 10 times (e.g., at least 15 times or even at least 20times) while retaining multipotency. Thus, culturing the ADSCs, withoutinducing differentiation, can be accomplished without specially screenedlots of serum. In contrast, mesenchymal stem cells (e.g., derived frombone marrow) would differentiate under the same culturing conditionsdescribed above. Methods for measuring viability and yield are known inthe art and can be employed (e.g., trypan blue exclusion).

[0085] The ADSCs can be separated by phenotypic identification, toidentify those cells that have two or more of the aforementioneddevelopmental lineages. To phenotypically separate the ADSCs from theADSC-EF, the cells are plated at a desired density, such as betweenabout 100 cells/cm² to about 100,000 cells/cm² (such as about 500cells/cm² to about 50,000 cells/cm², or, more particularly, betweenabout 1,000 cells/cm² to about 20,000 cells/cm²).

[0086] In a preferred embodiment the ADSC-EF is plated at a lowerdensity (e.g., about 300 cells/cm²) to facilitate the clonal isolationof the ADSCs. For example, after a few days, ADSCs plated at suchdensities will proliferate (expand) into a clonal population of ADSCs.

[0087] Such ADSCs can be used to clone and expand a multipotent ADSCinto clonal populations, using a suitable method for cloning cellpopulations. The cloning and expanding methods include cultures ofcells, or small aggregates of cells, physically picking and seeding intoa separate plate (such as the well of a multi-well plate).Alternatively, the stem cells can be subcloned onto a multi-well plateat a statistical ratio for facilitating placing a single cell into eachwell (e.g., from about 0.1 to about 1 cell/well or even about 0.25 toabout 0.5 cells/well, such as 0.5 cells/well). The ADSCs can be clonedby plating them at low density (e.g., in a petri-dish or other suitablesubstrate) and isolating them from other cells using devices such as acloning rings. Alternatively, where an irradiation source is available,clones can be obtained by permitting the cells to grow into a monolayerand then shielding one and irradiating the rest of cells within themonolayer. The surviving cell then will grow into a clonal population.While production of a clonal population can be expanded in any suitableculture medium, a preferred culture condition for cloning stem cells(such as the inventive stem cells or other stem cells) is about ⅔ F₁₂medium+20% serum (preferably fetal bovine serum) and about ⅓ standardmedium that haw been conditioned with stromal cells (e.g., cells fromthe stromal vascular fraction of liposuction aspirate), the relativeproportions being determined volumetrically).

[0088] In any event, whether clonal or not, the isolated ADSCs can becultured in a specific inducing medium to induce the ADSC todifferentiate and express its multipotency. The ADSCs give rise to cellsof mesodermal, ectodermal and endodermal lineage, and combinationsthereof. Thus, one or more ADSCs derived from a multipotent ADSC can betreated to differentiate into a variety of cell types.

[0089] In another embodiment, the ADSCs are cultured in a defmed mediumfor inducing adipogenic differentiation. Examples of specifc media thatinduce the ADSCs of the invention to take on a adipogenic phenotypeinclude, but are not limited to media containing a glucocorticoid (e.g.,dexamethasone, hydrocortisone, cortisone, etc.), insulin, a compoundwhich elevates intracellular levels of cAMP (e.g., dibutyryl-cAMP,8-CPT-cAMP (8-(4)chlorophenylthio)-adenosine 3′,5′ cyclic monophosphate;8-bromo-cAMP; dioctanoyl-cAMP, forskolin etc.), and/or a compound whichinhibits degradation of cAMP (e.g., a phosphodiesterase inhibitor suchas isobutyl methyl xanthine (IBMX), methyl isobutylxanthine,theophylline, caffeine, indomethacin, and the like), and serum. Thus,exposure of the ADSCs to between about 1 μM and about 10 μM insulin incombination with about 10⁻⁹ M to about 10⁻⁶ M to (e.g., about 1 μM)dexamethasone can induce adipogenic differentiation. Such a medium alsocan include other agents, such as indomethacin (e.g., about 100 μM toabout 200 μM), if desired, and preferably the medium is serum-free.

[0090] In another embodiment, ADSCs cultured in DMEM, 10% FBS, 1 uMdexamthasone, 10 uM insulin, 200 uM indomethacin, 1%antibiotic/antimicotic,(ABAM), 0.5 mM IBMX, take on an adipogenicphenotype.

[0091] Culturing media that can induce osteogenic differentiation of theADSCs include, but are not limited to, about 10⁻⁷ M and about 10⁻⁹ Mdexamethasone (e.g., about 1 μM) in combination with about 10 μM toabout 50 μM ascorbate-2-phosphate and between about 10 nM and about 50nM β-glycerophosphate. The medium also can include serum (e.g., bovineserum, horse serum, etc.).

[0092] In another embodiment, ADSCs cultured in DMEM, 10%FBS, 5% horseserum, 50 μM hydrocortisone, 10⁻⁷M dexamethosone, 50Mascorbate-2-phosphate, 1% ABAM, take on an osteogenic phenotype.

[0093] Culturing medium that can induce myogenic differentiation of theADSCs of the invention include, but is not limited to, exposing thecells to between about 10 μM and about 100 μM hydrocortisone, preferablyin a serum-rich medium (e.g., containing between about 10% and about 20%serum (either bovine, horse, or a mixture thereof)). Otherglucocorticoids that can be used include, but are not limited to,dexamethasone. Alternatively, 5′-azacytidine can be used instead of aglucocorticoid.

[0094] In another embodiment, ADSCs cultured in DMEM, 10% FBS, 10⁻⁷Mdexamethosone, 50 uMascorbate-2-phosphate, 10 mMbeta-glycerophosphate,1% ABAM, take on an myogenic phenotype.

[0095] Culturing medium that can induce chondrogenic differentiation ofthe ADSCs of the invention, include but is not limited to, exposing thecells to between about 1 μM to about 10 μM insulin and between about 1μM to about 10 μM transferrin, between about 1 ng/ml and 10 ng/mltransforming growth factor (TGF) β1, and between about 10 nM and about50 nM ascorbate-2-phosphate (50 nM). For chondrogenic differentiation,preferably the cells are cultured in high density (e.g., at aboutseveral million cells/ml or using micromass culture techniques), andalso in the presence of low amounts of serum (e.g., from about 1% toabout 5%).

[0096] In another embodiment, ADSCs cultured in DMEM, 50uMascorbate-2-phosphate, 6.25 ug/ml transferrin, 100 ng/ml insulingrowth factor (IGF-1), 5 ng/ml TGF-beta-1, 5 ng/ml basic fibroblastgrowth factor (bFGF; used only for one week), assume an chondrogenicphenotype.

[0097] In yet another embodiment, ADSCs are cultured in a neurogenicmedium such as DMEM, no serum and 5-10 mM β-mercaptoethanol and assumean ectodernal lineage.

[0098] The ADSCs also can be induced to dedifferentiate into adevelopmentally more immature phenotype (e.g., a fetal or embryonicphenotype). Such an induction is achieved upon exposure of the ADSC toconditions that mimic those within fetuses and embryos. For example, theinventive ADSCs, or population of ADSCs, can be co-cultured with cellsisolated from fetuses or embryos, or in the presence of fetal serum.

[0099] The ADSCs of the invention can be induced to differentiate into amesodermal, ectodermal, or an endodermal lineage by co-culturing theADSCs with mature cells from the respective germ layer, or precursorsthereof.

[0100] In an embodiment, induction of the ADSCs into specific cell typesby co-culturing with differentiated mature cells includes, but is notlimited to, myogenic differentiation induced by co-culturing the ADSCswith myocytes or myocyte precursors. Induction of the ADSCs into aneural lineage by co-culturing with neurons or neuronal precursors, andinduction of the ADSCs into an endodermal lineage, may occur byco-culturing with mature or precursor pancreatic cells or maturehepatocytes or their respective precursors.

[0101] Alternatively, the ADSCs are cultured in a conditioned medium andinduced to differentiate into a specific phenotype. Conditioned mediumis medium which was cultured with a mature cell that provides cellularfactors to the medium such as cytokines, growth factors, hormones, andextracellular matrix. For example, a medium that has been exposed tomature myoctytes is used to culture and induce ADSCs to differentiateinto a myogenic lineage. Other examples of conditioned media inducingspecific differentiation include, but are not limited to, culturing in amedium conditioned by exposure to heart valve cells to inducedifferentiation into heart valve tissue. In addition, ADSCs are culturedin a medium conditioned by neurons to induce a neuronal lineage, orconditioned by hepatoycytes to induce an endodermal lineage.

[0102] For co-culture, it may be desirable for the ADSCs and the desiredother cells to be co-cultured under conditions in which the two celltypes are in contact. This can be achieved, for example, by seeding thecells as a heterogeneous population of cells onto a suitable culturesubstrate. Alternatively, the ADSCs can first be grown to confluence,which will serve as a substrate for the second desired cells to becultured within the conditioned medium.

[0103] Other methods of inducing differentiation are known in the artand can be employed to induce the ADSCs to give rise to cells having amesodermal, ectodermal or endodermal lineage.

[0104] After culturing the stem cells in the differentiating-inducingmedium for a suitable time (e.g., several days to a week or more), theADSCs can be assayed to determine whether, in fact, they have acquiredthe desired lineage.

[0105] Methods to characterize differentiated cells that develop fromthe ADSCs of the invention, include, but are not limited to,histological, morphological, biochemical and immunohistochemicalmethods, or using cell surface markers, or genetically or molecularly,or by identifying factors secreted by the differentiated cell, and bythe inductive qualities of the differentiated ADSCs.

[0106] Molecular markers that characterize mesodermal cell thatdifferentiate from the ADSCs of the invention, include, but are notlimited to, MyoD, myosin, alpha-actin, brachyury, xFOG, Xtbx5 FoxF1,XNkx-2.5. Mammalian homologs of the above mentioned markers arepreferred.

[0107] Molecular markers that characterize ectodermal cell thatdifferentiate from the ADSCs of the invention, include but are notlimited to N-CAM, GABA and epidermis specific keratin. Mammalianhomologs of the above mentioned markers are preferred.

[0108] Molecular markers that characterize endodermal cell thatdifferentiate from the ADSCs include, but are not limited to, Xhbox8,Endo1, Xhex, Xcad2, Edd, EF1-alpha, HNF3-beta, LFABP, albumin, insulin.Mammalian homologs of the above mentioned markers are preferred.

[0109] In an embodiment, molecular characterization of thedifferentiated ADSCs is by measurement of telomere length.Undifferentiated stem cells have longer telomeres than differentiatedcells; thus the cells can be assayed for the level of telomeraseactivity. Alternatively, RNA or proteins can be extracted from the ADSCsand assayed (via Northern hybridization, RTPCR, Western blot analysis,etc.) for the presence of markers indicative of a specific phenotype.

[0110] In an alternative embodiment, differentiation is assessed byassaying the cells immunohistochemically or histologically , usingtissue-specific antibodies or stains, respectively. For example, toassess adipogenic differentiation, the differentiated ADSCs are stainedwith fat-specific stains (e.g., oil red O, safarin red, sudan black,etc.) or with labeled antibodies or molecular markers that identifyadipose-related factors (e.g., PPAR-γ, adipsin, lipoprotein lipase,etc.).

[0111] In another embodiment, ostogenesis can be assessed by stainingthe differentiated ADSCs with bone-specific stains (e.g., alkalinephosphatase, von Kossa, etc.) or with labeled antibodies or molecularmarkers that identify bone-specific markers (e.g., osteocalcin,osteonectin, osteopontin, type I collagen, bone morphogenic proteins,cbfa, etc.).

[0112] Myogensis can be assessed by identifying classical morphologicchanges (e.g., polynucleated cells, syncitia formation, etc.), orassessed biochemically for the presence of muscle-specific factors(e.g., myo D, myosin heavy chain, etc.).

[0113] Chondrogenesis can be determined by staining the cells usingcartilage-specific stains (e.g., Alcian blue) or with labeled antibodiesor molecular markers that identify cartilage-specific molecules (e.g.,sulfated glycosaminoglycans and proteoglycans, keratin, chondroitin,Type II collagen, etc.) in the medium.

[0114] Alternative embodiments can employ methods of assessingdevelopmental phenotype, known in the art. For example, the cells can besorted by size and granularity. The cells can be used as an immunogen togenerate monoclonal antibodies (Kohler and Milstein), which can then beused to bind to a given cell type. Correlation of antigenicity canconfirm that the ADSC has differentiated along a given developmentalpathway.

[0115] While an ADSC can be isolated, preferably it is within apopulation of cells. The invention provides a defmed population ofADSCs. In an embodiment, the population is heterogeneous. In anotherembodiment, the population is homogeneous.

[0116] In another embodiment, a population of ADSCs can support cellsfor culturing other cells. For example, cells that can be supported byADSC populations include other types of stem cells, such as neural stemcells (NSC), hematopoetic stem cells (HPC, particularly CD34⁺ stemcells), embryonic stem cells (ESC) and mixtures thereof). In otherembodiments, the population is substantially homogeneous, consistingessentially of the inventive adipose-derived stem cells.

Uses of the ADSC-EF, ADSCs and Methods of the Invention

[0117] The ADSC-EF can be used as a source of the ADSCs of theinvention. The ADSC-EF can be introduced into a subject for tissueregeneration, wound repair or other applications requiring a source ofstem cells. In addition, the ADSC-EF can be treated to cause the ADSCstherein to differentiate into a desired cell type for introduction intoa subject. The ADSC-EF can also be cultured in vitro to maintain asource of ADSCs, or can be induced to produce further differentiatedADSCs that can develop into a desired tissue.

[0118] The ADSCs (and populations of ADSCs) can be employed for avariety of purposes. The ADSCs can support the growth and expansion ofother cell types. The invention includes a method of conditioningculture medium using the ADSCs in a suitable medium, and theADSC-conditioned medium produced by such a method. Typically, the mediumis used to support the in vitro growth of the ADSCs, which secretehormones, cell matrix material, and other factors into the medium. Aftera suitable period (e.g., one or a few days), the culture mediumcontaining the secreted factors can be separated from the cells andstored for future use. The ADSCs can be re-used successively tocondition medium, as desired. In other applications (e.g., forco-culturing the ADSCs with other cell types), the cells can remainwithin the conditioned medium. Thus, the invention provides anADSC-conditioned medium obtained using this method, which either cancontain the ADSCs, or be substantially free of the ADSCs, as desired.

[0119] The ADSC-conditioned medium can be used to support the growth andexpansion of desired cell types, and the invention provides a method ofculturing cells (particularly stem cells) using the conditioned medium.The method involves maintaining a desired cell in the conditioned mediumunder conditions for the cell to remain viable. The cell can bemaintained under any suitable condition for culturing them, such as areknown in the art. Desirably, the method permits successive rounds ofmitotic division of the cell to form an expanded population. The exactconditions (e.g., temperature, CO₂ levels, agitation, presence ofantibiotics, etc.) will depend on the other constituents of the mediumand on the cell type. However, optimizing these parameters is within theordinary skill in the art.

[0120] In another embodiment, the ADSCs can be genetically modified,e.g., to express exogenous genes (“transgenes”) or to repress theexpression of endogenous genes, and the invention provides a method ofgenetically modifying such cells and populations. In accordance withthis method, the ADSC is exposed to a gene transfer vector comprising anucleic acid including a transgene, such that the nucleic acid isintroduced into the cell under conditions appropriate for the transgeneto be expressed within the cell. The transgene generally is anexpression cassette, including a polynucleotide operably linked to asuitable promoter. The polynucleotide can encode a protein, or it canencode biologically active RNA (e.g., antisense RNA or a ribozyme).Thus, for example, the polynucleotide can encode a gene conferringresistance to a toxin, a hormone (such as peptide growth hormones,hormone releasing factors, sex hormones, adrenocorticotrophic hormones,cytokines (e.g., interfering, interleukins, lymphokines), etc.), acell-surface-bound intracellular signaling moiety (e.g., cell adhesionmolecules, hormone receptors, etc.), a factor promoting a given lineageof differentiation, (e.g., bone morphogenic protein (BMP)) etc. Ofcourse, where it is desired to employ gene transfer technology todeliver a given transgene, its sequence will be known.

[0121] Within the expression cassette, the coding polynucleotide isoperably linked to a suitable promoter. Examples of suitable promotersinclude prokaryotic promoters and viral promoters (e.g., retroviralITRs, LTRs, immediate early viral promoters (IEp), such as herpesvirusIEp (e.g., ICP4-IEp and ICP0-IEp), cytomegalovirms (CMV) lEp, and otherviral promoters, such as Rous Sarcoma Virus (RSV) promoters, and MurineLeukemia Virus (MLV) promoters). Other suitable promoters are eukaryoticpromoters, such as enhancers (e.g., the rabbit β-globin regulatoryelements), constitutively active promoters (e.g., the β-actin promoter,etc.), signal specific promoters (e.g., inducible promoters such as apromoter responsive to RU486, etc.), and tissue-specific promoters. Itis well within the skill of the art to select a promoter suitable fordriving gene expression in a predefined cellular context. The expressioncassette can include more than one coding polynucleotide, and it caninclude other elements (e.g., polyadenylation sequences, sequencesencoding a membrane-insertion signal or a secretion leader, ribosomeentry sequences, transcriptional regulatory elements (e.g., enhancers,silencers, etc.), and the like), as desired.

[0122] The expression cassette containing the transgene should beincorporated into a genetic vector suitable for delivering the transgeneto the cells. Depending on the desired end application, any such vectorcan be so employed to genetically modify the cells (e.g., plasmids,naked DNA, viruses such as adenovirus, adeno-associated virus,herpesviruses, lentiviruses, papillomaviruses, retroviruses, etc.). Anymethod of constructing the desired expression cassette within suchvectors can be employed, many of which are well known in the art (e.g.,direct cloning, homologous recombination, etc.). Of course, the choiceof vector will largely determine the method used to introduce the vectorinto the cells (e.g., by protoplast fusion, calcium-phosphateprecipitation, gene gun, electroporation, infection with viral vectors,etc.), which are generally known in the art.

[0123] The genetically altered ADSCs can be employed as bioreactors forproducing the product of the transgene. In other embodiments, thegenetically modified ADSCs are employed to deliver the transgene and itsproduct to an animal. For example, the ADSCs, once genetically modified,can be introduced into the animal under conditions sufficient for thetransgene to be expressed in vivo.

[0124] In addition to serving as useful targets for geneticmodification, many ADSCs and populations of ADSCs secrete hormones(e.g., cytokines, peptide or other (e.g., monobutyrin) growth factors,etc.). Some of the cells naturally secrete such hormones upon initialisolation, and other cells can be genetically modified to secretehormones, as discussed herein. The ADSCs that secrete hormones can beused in a variety of contexts in vivo and in vitro. For example, suchcells can be employed as bioreactors to provide a ready source of agiven hormone, and the invention pertains to a method of obtaininghormones from such cells. In accordance with the method, the ADSCs arecultured, under suitable conditions for them to secrete the hormone intothe culture medium. After a suitable period of time, and preferablyperiodically, the medium is harvested and processed to isolate thehormone from the medium. Any standard method (e.g., gel or affinitychromatography, dialysis, lyophilization, etc.) can be used to purifythe hormone from the medium, many of which are known in the art.

[0125] In other embodiments, ADSCs (and populations) secreting hormonescan be employed as therapeutic agents. Generally, such methods involvetransferring the cells to desired tissue, either in vitro (e.g., as agraft prior to implantation or engrafting) or in vivo, to animal tissuedirectly. The cells can be transferred to the desired tissue by anymethod appropriate, which generally will vary according to the tissuetype. For example, ADSCs can be transferred to a graft by bathing thegraft (or infusing it) with culture medium containing the cells.Alternatively, the ADSCs can be seeded onto the desired site within thetissue to establish a population. Cells can be transferred to sites invivo using devices such as catheters, trocars, cannulae, stents (whichcan be seeded with the cells), etc. For these applications, preferablythe ADSC secretes a cytokine or growth hormone such as human growthfactor, fibroblast growth factor, nerve growth factor, insulin-likegrowth factors, hemopoietic stem cell growth factors, members of thefibroblast growth factor family, members of the platelet-derived growthfactor family, vascular and endothelial cell growth factors, members ofthe TGFb family (including bone morphogenic factor), or enzymes specificfor congenital disorders (e.g., dystrophic).

[0126] In one application, the invention provides a method of promotingthe closure of a wound within a patient using ADSCs. In accordance withthe method, ADSCs secreting the hormone are transferred to the vicinityof a wound under conditions sufficient for the cells to produce thehormone. The presence of the hormone in the vicinity of the woundpromotes closure of the wound. The method promotes closure of bothexternal (e.g., surface) and internal wounds. Wounds to which thepresent inventive method is useful in promoting closure include, but arenot limited to, abrasions, avulsions, blowing wounds, bum wounds,contusions, gunshot wounds, incised wounds, open wounds, penetratingwounds, perforating wounds, puncture wounds, seton wounds, stab wounds,surgical wounds, subcutaneous wounds, or tangential wounds. The methodneed not achieve complete healing or closure of the wound; it issufficient for the method to promote any degree of wound closure. Inthis respect, the method can be employed alone or as an adjunct to othermethods for healing wounded tissue.

[0127] Where the ADSCs secrete an angiogenic hormone (e.g., vasculargrowth factor, vascular and endothelial cell growth factor, etc.), they(as well as populations containing them) can be employed to induceangiogenesis within tissues. Thus, the invention provides a method ofpromoting or inhibiting neovascularization within tissue using suchADSCs. The presence of the hormone within the tissue promotes orinhibits neovascularization. In accordance with this method, the ADSC isintroduced the desired tissue under conditions sufficient for the cellto produce the angiogenic hormone. The presence of the hormone withinthe tissue promotes neovascularization within the tissue.

[0128] Because the ADSCs have a developmental phenotype, they can beemployed in tissue engineering. In this regard, the invention provides amethod of producing animal matter comprising maintaining the ADSCs underconditions sufficient for them to expand and differentiate to form thedesired matter. The matter can include mature tissues, or even wholeorgans, including tissue types into which the inventive cells candifferentiate (as set forth herein). Typically, such matter willcomprise adipose, cartilage, heart, dermal connective tissue, bloodtissue, muscle, kidney, bone, pleural, splanchnic tissues, vasculartissues, and the like. More typically, the matter will comprisecombinations of these tissue types (i.e., more than one tissue type).For example, the matter can comprise all or a portion of an animal organ(e.g., a heart, a kidney) or a limb (e.g., a leg, a wing, an arm, ahand, a foot, etc.). Of course, in as much as the cells can divide anddifferentiate to produce such structures, they can also form anlagen ofsuch structures. At early stages, such anlagen can be cryopreserved forfuture generation of the desired mature structure or organ.

[0129] To produce such structures, the ADSCs and populations aremaintained under conditions suitable for them to expand and divide toform the desired structures. In some applications, this is accomplishedby transferring them to an animal (i.e., in vivo) typically at a sightat which the new matter is desired. Thus, for example, the invention canfacilitate the regeneration of tissues (e.g., bone, muscle, cartilage,tendons, adipose, etc.) within an animal where the ADSCs are implantedinto such tissues. In other embodiments, and particularly to createanlagen, the ADSCs can be induced to differentiate and expand intotissues in vitro. In such applications, the ADSCs are cultured onsubstrates that facilitate formation into three-dimensional structuresconducive for tissue development. Thus, for example, the ADSCs can becultured or seeded onto a bio-compatible lattice, such as one thatincludes extracellular matrix material, synthetic polymers, cytokines,growth factors, etc. Such a lattice can be molded into desired shapesfor facilitating the development of tissue types. Also, at least at anearly stage during such culturing, the medium and/or substrate issupplemented with factors (e.g., growth factors, cytokines,extracellular matrix material, etc.) that facilitate the development ofappropriate tissue types and structures. Indeed, in some embodiments, itis desired to co-culture the ADSCs with mature cells of the respectivetissue type, or precursors thereof, or to expose the cells to therespective conditioned medium, as discussed herein.

[0130] To facilitate the use of the ADSCs and populations for producingsuch animal matter and tissues, the invention provides a compositionincluding the ADSCs (and populations) and biologically compatiblelattice. Typically, the lattice is formed from polymeric material,having fibers as a mesh or sponge, typically with spaces on the order ofbetween about 100 μm and about 300 μm. Such a structure providessufficient area on which the cells can grow and proliferate. Desirably,the lattice is biodegradable over time, so that it will be absorbed intothe animal matter as it develops. Suitable polymeric lattices, thus, canbe formed from monomers such as glycolic acid, lactic acid, propylfumarate, caprolactone, hyaluronan, hyaluronic acid, and the like. Otherlattices can include proteins, polysaccharides, polyhydroxy acids,polyorthoesters, polyanhydrides, polyphosphazenes, or synthetic polymers(particularly biodegradable polymers). Of course, a suitable polymer forforming such lattice can include more than one monomers (e.g.,combinations of the indicated monomers). Also, the lattice can alsoinclude hormones, such as growth factors, cytokines, and morphogens(e.g., retinoic acid, aracadonic acid, etc.), desired extracellularmatrix molecules (e.g., fibronectin, laminin, collagen, etc.), or othermaterials (e.g., DNA, viruses, other cell types, etc.) as desired.

[0131] To form the composition, the ADSCs are introduced into thelattice such that they permeate into the interstitial spaces therein.For example, the matrix can be soaked in a solution or suspensioncontaining the cells, or they can be infused or injected into thematrix. A particularly preferred composition is a hydrogel formed bycrosslinking of a suspension including the polymer and also having theinventive cells dispersed therein. This method of formation permits thecells to be dispersed throughout the lattice, facilitating more evenpermeation of the lattice with the cells. Of course, the compositionalso can include mature cells of a desired phenotype or precursorsthereof, particularly to potentate the induction of the ADSCs todifferentiate appropriately within the lattice (e.g., as an effect ofco-culturing such cells within the lattice).

[0132] The composition can be employed in any suitable manner tofacilitate the growth and generation of the desired tissue types,structures, or anlagen. For example, the composition can be constructedusing three-dimensional or sterotactic modeling techniques. Thus, forexample, a layer or domain within the composition can be populated bycells primed for osteogenic differentiation, and another layer or domainwithin the composition can be populated with cells primed for myogenicand/or chondrogenic development. Bringing such domains intojuxtaposition with each other facilitates the molding anddifferentiation of complex structures including multiple tissue types(e.g., bone surrounded by muscle, such as found in a limb). To directthe growth and differentiation of the desired structure, the compositioncan be cultured ex vivo in a bioreactor or incubator, as appropriate. Inother embodiments, the structure is implanted within the host animaldirectly at the site in which it is desired to grow the tissue orstructure. In still another embodiment, the composition can be engraftedon a host (typically an animal such as a pig, baboon, etc.), where itwill grow and mature until ready for use. Thereafter, the maturestructure (or anlagen) is excised from the host and implanted into thehost, as appropriate.

[0133] Lattices suitable for inclusion into the composition can bederived from any suitable source (e.g., matrigel), and some commercialsources for suitable lattices exist (e.g., suitable of polyglycolic acidcan be obtained from sources such as Ethicon, N.J.). Another suitablelattice can be derived from the acellular portion of adiposetissue—i.e., adipose tissue extracellular matrix matter substantiallydevoid of cells, and the invention provides such a adipose-derivedlattice. Typically, such adipose-derived lattice includes proteins suchas proteoglycans, glycoproteins, hyaluronins, fibronectins, collagens(type I, type II, type III, type IV, type V, type VI, etc.), and thelike, which serve as excellent substrates for cell growth. Additionally,such adipose-derived lattices can include hormones, preferably cytokinesand growth factors, for facilitating the growth of cells seeded into thematrix.

[0134] The adipose-derived matrix can be isolated form adipose tissuesimilarly as described above, except that it will be present in theacellular fraction. For example, adipose tissue or derivatives thereof(e.g; the lattice enriched supernant fraction of the method describedabove) can be subjected to sonic or thermal energy and/or enzymaticprocessed to recover the matrix material. Also, desirably the cellularfraction of the adipose tissue is disrupted, for example by treating itwith lipases, detergents, proteases, and/or by mechanical or sonicdisruption (e.g., using a homogenizer or sonicator). However isolated,the material is initially identified as a viscous material, but it canbe subsequently treated, as desired, depending on the desired end use.For example, the raw matrix material can be treated (e.g., dialyzed ortreated with proteases or acids, etc.) to produce a desirable latticematerial. Thus the lattice can be prepared in a hyrated form or it canbe dried or lyophilized into a substantially anhydrous form or a powder.Thereafter, the powder can be rehydrated for use as a cell culturesubstrate, for example by suspending it in a suitable cell culturemedium. In this regard, the adipose-derived lattice can be mixed withother suitable lattice materials, such as described above. Of course,the invention pertains to compositions including the adipose-derivedlattice and cells or populations of cells, such as the inventive ADSCsand other cells as well (particularly other types of stem cells).

[0135] As discussed above, the ADSCs, populations, lattices, andcompositions of the invention can be used in tissue engineering andregeneration. Thus, the invention pertains to an implantable structure(i.e., an implant) incorporating any of these inventive features. Theexact nature of the implant will vary according to the use to which itis to be put. The implant can be or comprise, as described, maturetissue, or it can include immature tissue or the lattice. Thus, forexample, one type of implant can be a bone implant, comprising apopulation of the inventive cells that are undergoing (or are primedfor) osteogenic differentiation, optionally seeded within a lattice of asuitable size and dimension, as described above. Such an implant can beinjected or engrafted within a host to encourage the generation orregeneration of mature bone tissue within the patient. Similar implantscan be used to encourage the growth or regeneration of muscle, fat,cartilage, tendons, etc., within patients. Other types of implants areanlagen (such as described herein), e.g., limb buds, digit buds,developing kidneys, etc, that, once engrafted onto a patient, willmature into the appropriate structures.

[0136] The adipose-derived lattice can conveniently be employed as partof a cell culture kit. Accordingly, the invention provides a kitincluding the inventive adipose-derived lattice and one or more othercomponents, such as hydrating agents (e.g., water,physiologically-compatible saline solutions, prepared cell culturemedia, serum or derivatives thereof, etc.), cell culture substrates(e.g., culture dishes, plates, vials, etc.), cell culture media (whetherin liquid or powdered form), antibiotic compounds, hormones, and thelike. While the kit can include any such ingredients, preferably itincludes all ingredients necessary to support the culture and growth ofdesired cell types upon proper combination. Of course, if desired, thekit also can include cells (typically frozen), which can be seeded intothe lattice as described herein.

[0137] While many aspects of the invention pertain to tissue growth anddifferentiation, the invention has other applications as well. Forexample, the adipose-derived lattice can be used as an experimentalreagent, such as in developing improved lattices and substrates fortissue growth and differentiation. The adipose-derived lattice also canbe employed cosmetically, for example, to hide wrinkles, scars,cutaneous depressions, etc., or for tissue augmentation. For suchapplications, preferably the lattice is stylized and packaged in unitdosage form. If desired, it can be admixed with carriers (e.g., solventssuch as glycerine or alcohols), perfumes, antibiotics, colorants, andother ingredients commonly employed in cosmetic products. The substratealso can be employed autologously or as an allograft, and it can usedas, or included within, ointments or dressings for facilitating woundhealing. The ADSCs can also be used as experimental reagents. Forexample, they can be employed to help discover agents responsible forearly events in differentiation. For example, the inventive cells can beexposed to a medium for inducing a particular line of differentiationand then assayed for differential expression of genes (e.g., byrandom-primed PCR or electrophoresis or protein or RNA, etc.).

[0138] As any of the steps for isolating the inventive ADSCs or theadipose-derived lattice, the, the invention provides a kit for isolatingsuch reagents from adipose tissues. The kit can include a means forisolating adipose tissue from a patient (e.g., a cannula, a needle, anaspirator, etc.), as well as a means for separating stromal cells (e.g.,through methods described herein). The kit can be employed, for example,as an immediate source of ADSCs that can then be re-introduced from thesame individual as appropriate. Thus, the kit can facilitate theisolation of adipose-derived stem cells for implantation in a patientneeding regrowth of a desired tissue type, even in the same procedure.In this respect, the kit can also include a medium for differentiatingthe cells, such as those set forth herein. As appropriate, the cells canbe exposed to the medium to prime them for differentiation within thepatient as needed. In addition, the kit can be used as a convenientsource of ADSCs for in vitro manipulation (e.g., cloning ordifferentiating as described herein). In another embodiment, the kit canbe employed for isolating a adipose-derived lattice as described herein.

[0139] While one of skill in the art is fully able to practice theinstant invention upon reading the foregoing detailed description, thefollowing examples will help elucidate some of its features. Inparticular, they demonstrate the isolation of a human adipose-derivedstem cell substantially free of mature adipocytes, the isolation of aclonal population of such cells, the ability of such cells todifferentiate in vivo and in vitro into all cells with a mesodermalphenotype, endodermal phenotype, and extodermal phenotype, and thecapacity of such cells to support the growth of other types of stemcells. The examples also demonstrate the isolation of a adipose-derivedlattice substantially free of cells that is capable of serving as asuitable substrate for cell culture. Of course, as these examples arepresented for purely illustrative purposes, they should not be used toconstrue the scope of the invention in a limited manner, but rathershould be seen as expanding upon the foregoing description of theinvention as a whole.

[0140] The procedures employed in these examples, such as surgery, cellculture, enzymatic digestion, histology, and molecular analysis ofproteins and polynucleotides, are familiar to those of ordinary skill inthis art. As such, and in the interest of brevity, experimental detailsare not recited in detail.

EXAMPLE 1

[0141] This example demonstrates the isolation of a humanadipose-derived stem cell substantially free of mature adipocytes.

[0142] Raw liposuction aspirate was obtained from patients undergoingelective surgery. Prior to the liposuction procedures, the patients weregiven epinephrine to minimize contamination of the aspirate with blood.The aspirate was strained to separate associated adipose tissue piecesfrom associated liquid waste. Isolated tissue was rinsed thoroughly withneutral phosphate buffered saline and then enzymatically dissociatedwith 0.075% w/v collagenase at 37° C. for about 20 minutes underintermittent agitation. Following the digestion, the collagenase wasneutralized, and the slurry was centrifuged at about 260 g for about 10minutes, which produced a multi-layered supernatant and a cellularpellet. The supernatant was removed and retained for further use, andthe pellet was resuspended in an erythrocyte-lysing solution andincubated without agitation at about 25° C. for about 10 minutes.Following incubation, the medium was neutralized, and the cells wereagain centrifuged at about 250 g for about 10 minutes. Following thesecond centrifugation, the cells were suspended, and assessed forviability (using trypan blue exclusion) and cell number. Thereafter,they were plated at a density of about at about 1×10⁶ cells/100 mm dish.They were cultured at 37° C. in DMEM+fetal bovine serum (about 10%) inabout 5% CO₂.

[0143] The majority of the cells were adherent, small, mononucleic,relatively agranular fibroblast-like cells containing no visible lipiddroplets and were CD34-negative. The majority of the cells stainednegatively with oil-red O and von Kossa. The cells were also assayed forexpression of telomerase (using a commercially available TRAP assaykit), using HeLa cells and HN-12 cells as positive controls. Humanforeskin fibroblasts and HN-12 heated cell extracts were used asnegative controls. Telomeric products were resolved onto a 12.5%polyacrylamide cells and the signals determined by phosphorimaging.Telemeric ladders representing telomerase activity were observed in theadipose-derived stem cells as well as the positive controls. No ladderswere observed in the negative controls.

[0144] Thus, these cells were not identifiable as myocytes, adipocytes,chondrocytes, osteocytes, or blood cells. These results demonstrate thatthe adipose-derived cells express telomerase activity similar to thatpreviously reported for human stem cells.

[0145] Subpopulations of these cells were then exposed to the followingmedia to assess their developmental phenotype: Adipogenesis OsteogenesisMyogenesis Chondrogenesis DMEM DMEM DMEM DMEM 10% FETAL 10% FETAL 10%FETAL 1% FETAL BOVINE BOVINE BOVINE BOVINE SERUM SERUM SERUM SERUM 0.5mM ISO-  5% horse serum  5% horse serum 6.25 μg/ml insulin BUTYL- 0.1 μM50 μM 6.25 μg/ml METHYL- dexamethasone hydrocortisone transferrinXANTHINE  50 μM ascorbate-  1% ABAM   10 ng/ml  1 μM 2-phosphate TGFβ1dexamethasone  10 mM β-   50 nM  10 μM insulin glycerophosphateascorbate-2- 200 μM  1% ABAM phosphate indomethacin 1% ABAM  1% ABAM

[0146] A population was cultured at high density in the chondrogenicmedium for several weeks. Histological analysis of the tissue cultureand paraffin sections was performed with H&E, alcian blue, toludeneblue, and Goldner's trichrome staining at 2, 7, and 14 days.Immunohistochemistry was performed using antibodies againstchondroitin-4-sulfate and keratin sulfate and type II collagen.Qualitative estimate of matrix staining was also performed. The resultsindicated that cartilaginous spheroid nodules with a distinct border ofperichondral cells formed as early as 48 hours after initial treatment.Untreated control cells exhibited no evidence of chondrogenicdifferentiation. These results confirm that the stem cells havechondrogenic developmental phenotype.

[0147] A population was cultured until near confluence and then exposedto the adipogenic medium for several weeks. The population was examinedat two and four weeks after plating by calorimetric assessment ofrelative opacity following oil red-O staining. Adipogenesis wasdetermined to be underway at two weeks and quite advanced at four weeks(relative opacity of 1 and 5.3, respectively). Bone marrow-derived stemcells were employed as a positive control, and these cells exhibitedslightly less adipogenic potential (relative density of 0.7 and 2.8,respectively).

[0148] A population was cultured until near confluence and then exposedto the osteogeneic medium for several weeks. The population was examinedat two and four weeks after plating by colorimetric assessment ofrelative opacity following von Kossa staining. Osteogenesis wasdetermined to be underway at two weeks and quite advanced at four weeks(relative opacity of 1.1 and 7.3, respectively. Bone marrow-derived stemcells were employed as a positive control, and these cells exhibitedslightly less osteogenic potential (relative density of 0.2 and 6.6,respectively).

[0149] A population was cultured until near confluence and then exposedto the myogenic medium for several weeks. The population was examined atone, three, and six weeks after plating by assessment of multinucleatedcells and expression of muscle-specific proteins (MyoD and myosin heavychain). Human foreskin fibroblasts and skeletal myoblasts were used ascontrols. Cells expressing MyoD and myosin were found at all time pointsfollowing exposure to the myogenic medium in the stem cell population,and the proportion of such cells increased at 3 and 6 weeks.Multinucleated cells were observed at 6 weeks. In contrast, thefibroblasts exhibited none of these characteristics at any time points.

[0150] These results demonstrate the isolation of a humanadipose-derived pluripotent stem cell substantially free of matureadipocytes.

EXAMPLE 2

[0151] This example demonstrates that the adipose-derived stem cells donot differentiate in response to 5-azacytidine.

[0152] Adipose-derived stem cells obtained in accordance with Example 1were cultured in the presence of 5-azacytidine. In contrast with bonemarrow-derived stem cells, exposure to this agent did not inducemyogenic differentiation (see Wakitani et al., supra).

EXAMPLE 3

[0153] This example demonstrates the generation of a clonal populationof human adipose-derived stem cells from an adipose-derived stem cellenriched fraction.

[0154] Cells isolated in accordance with the procedure set forth inExample 1 were plated at about 5,000 cells/100 mm dish and cultured fora few days as indicated in Example 1. After some rounds of celldivision, some clones were picked with a cloning ring and transferred towells in a 48 well plate. These cells were cultured for several weeks,changing the medium twice weekly, until they were about 80% to about 90%confluent (at 37° C. in about 5% CO₂ in ⅔ F₁₂ medium+20% fetal bovineserum and ⅓ standard medium that was first conditioned by the cellsisolated in Example 1, “cloning medium”). Thereafter, each culture wastransferred to a 35 mm dish and grown, and then retransferred to a 100mm dish and grown until close to confluent. Following this, one cellpopulation was frozen, and the remaining populations were plated on 12well plates, at 1000 cells/well.

[0155] The cells were cultured for more than 15 passages in cloningmedium and monitored for differentiation as indicated in Example 1. Theundifferentiated state of each clone remained true after successiverounds of culturing.

[0156] Populations of the clones then were established and exposed toadipogenic, chondrogenic, myogenic, and osteogenic medium as discussedin Example 1. It was observed that at least one of the clones was ableto differentiate into bone, fat, cartilage, and muscle when exposed tothe respective media, and most of the clones were able to differentiateinto at least three types of tissues. The capacity of the cells todevelop into muscle and cartilage further demonstrates thepluripotentiality of these adipose-derived stem cells.

[0157] These results demonstrate that the adipose-derived stem cells canbe maintained in an undifferentiated state for many passages without therequirement for specially pre-screened lots of serum. The results alsodemonstrate that the cells retain pluripotentiality following suchextensive passaging, proving that the cells are indeed stem cells andnot merely committed progenitor cells.

EXAMPLE 4

[0158] This example demonstrates the adipose-derived stem cells from anadipose-devired stem cell enriched fraction can support the culture ofother types of stem cells.

[0159] Human adipose-derived stem cells were passaged onto 96 wellplates at a density of about 30,000/well, cultured for one week and thenirradiated. Human CD34⁺ hematopoetic stem cells isolated from umbilicalcord blood were then seeded into the wells. Co-cultures were maintainedin MyeloCult H5100 media, and cell viability and proliferation weremonitored subjectively by microscopic observation. After two weeks ofco-culture, the hematopoetic stem cells were evaluated for CD34expression by flow cytometry.

[0160] Over a two-week period of co-culture with stromal cells, thehematopoetic stem cells formed large colonies of rounded cells. Flowanalysis revealed that 62% of the cells remained CD34⁺. Based onmicroscopic observations, human adipo-derived stromal cells maintainedthe survival and supported the growth of human hematopoetic stem cellsderived from umbilical cord blood.

[0161] These results demonstrate that stromal cells from humansubcutaneous adipose tissue are able to support the ex vivo maintenance,growth and differentiation of other stem cells.

EXAMPLE 5

[0162] This example demonstrates that the adipose-derived stem cellsfrom an adipose-devired stem cell enriched fraction can differentiate invivo.

[0163] Four groups (A-D) of 12 athymic mice each were implantedsubcutaneously with hydroxyapatite/tricalcium phosphate cubes containingthe following: Group A contained adipose-derived stem cells that hadbeen pretreated with osteogenic medium as set forth in Example 1. GroupB contained untreated adipose-derived stem cells. Group C containedosteogenic medium but no cells. Group D contained non-osteogenic mediumand no cells. Within each group, six mice were sacrificed at threeweeks, and the remaining mice sacrificed at eight weeks followingimplantation. The cubes were extracted, fixed, decalcified, andsectioned. Each section was analyzed by staining with hematoxylin andeosin (e.g., H&E), Mallory bone stain, and immunostaining forosteocalcin.

[0164] Distinct regions of osteoid-like tissue staining for osteocalcinand Mallory bone staining was observed in sections from groups A and B.Substantially more osteoid tissue was observed in groups A and B than inthe other groups (p<0.05 ANOVA), but no significant difference inosteogenesis was observed between groups A and B. Moreover, aqualitative increase in bone growth was noted in both groups A and Bbetween 3 and 8 weeks. These results demonstrate that theadipose-derived stem cells can differentiate in vivo.

EXAMPLE 6

[0165] This example demonstrates the isolation of an adipose-derivedlattice substantially devoid of cells.

[0166] In one protocol, the lattice-enriched fraction from Example 1 wassubjected to enzymatic digestion for three days in 0.05% trypsinEDTA/100 U/ml deoxyribonuclease to destroy the cells. Every day thedebris was rinsed in saline and fresh enzyme was added. Thereafter thematerial was rinsed in saline and resuspended in 0.05% collagease andabout 0.1% lipase to partially digest the proteins and fat present. Thisincubation continued for two days.

[0167] In another protocol, the withheld supernatant from Example 1 wasincubated in EDTA to eliminate any epithelial cells. The remaining cellswere lysed using a buffer containing 1% NP40, 0.5% sodium deoxycholate,0.1% SDS, 5 mM EDTA, 0.4M NaCl, 50 mMTris-HCL (pH 8) and proteaseinhibitors, and 10 μg/ml each of leupeptin, chymostatin, antipain, andpepstatin A. Finally, the tissue was extensively washed in PBS withoutdivalent cations.

[0168] After both preparatory protocols, remaining substance was washedand identified as a gelatinous mass. Microscopic analysis of thismaterial revealed that it contained no cells, and it was composed ofhigh amounts of collagen (likely type IV) and a wide variety of growthfactors. Preparations of this material have supported the growth ofcells, demonstrating it to be an excellent substrate for tissue culture.

EXAMPLE 7

[0169] The following description provides adipose-derived stem cellsenriched fractions which exhibit mesodermal multi-tissue potential, andmethods for isolating said stem cells.

Materials and Methods

[0170] All materials were purchased from Sigma (St. Louis, Mo.) unlessotherwise stated. All tissue culture reagents were purchased from LifeTechnologies (New York, N.Y.). Fetal Bovine Serum (FBS) and Horse Serum(HS) were purchased from Hyclone (Logan, Utah) and Life Technologies,respectively.

Cell Lines

[0171] Normal Human Osteoblasts (NHOsts), human Skeletal Muscle (SkM)cells and a population of Mesenchymal Stem Cells derived from bonemarrow (MSCs) were purchased from Clonetics (Walkersville, Md.). Themurine 3T3-L1 pre-adipocyte cell line (Green H., and Meuth, M., 1974,Cell 3: 127-133) was obtained from ATCC (Rockville, Md.). Human ForeskinFibroblasts (HFFs) were obtained from Cascade Biologics (Portland,Oreg.).

Isolation and Culture of Stem Cells

[0172] Human adipose tissue was obtained from elective liposuctionprocedures under local anesthesia according to patient consent protocol,HSPC #98-08 011-02 (Univerisity of California Los Angeles). In thisprocedure, a hollow blunt-tipped cannula was introduced into thesubcutaneous space through small (˜1 cm) incisions. The cannula wasattached to a gentle suction and moved through the adipose compartment,mechanically disrupting the fat tissue. A solution of saline and thevasoconstrictor, epinephrine, was infused into the adipose compartmentto minimize blood loss and contamination of the tissue by peripheralblood cells. The raw lipoaspirate (approximately 300 cc) was processedaccording to established methodologies in order to obtain a stromalvascular fraction (SVF) (Hauner H., et al., 1987, J. Clin. Endocrinol.Metabol. 64: 832-835; Katz, A. J., et al., 1999 Clin. Plast. Surg. 26:587-603, viii). To isolate the SVF, lipoaspirates were washedextensively with equal volumes of Phospho-Buffered Saline (PBS) and theextracellular matrix (ECM) was digested at 37° C. for 30 minutes with0.075% collagenase. Enzyme activity was neutralized with Dulbecco'sModified Eagle's Medium (DMEM), containing 10% Fetal Bovine Serum (FBS)and centrifuged at 1200×g for 10 minutes to obtain a high-density SVFpellet. The pellet was resuspended in 160 mM NH₄Cl and incubated at roomtemperature for 10 minutes to lyse contaminating red blood cells. TheSVF was collected by centrifugation, as detailed above, filtered througha 100 μm nylon mesh to remove cellular debris and incubated overnight at37° C./5% CO₂ in Control Medium (DMEM, 10% FBS, 1%antibiotic/antimycotic solution). Following incubation, the plates werewashed extensively with PBS to remove residual non-adherent red bloodcells. The resulting cell population was termed an adipose-derived stemcell enriched fraction (ADSC enriched fraction), in order to distinguishit from the SVF obtained from excised adipose tissue. Theadipose-derived stem cells were maintained at 37° C./5% CO₂ innon-inductive Control Medium. Cells did not require specific FBS seralots for expansion and differentiation . For immunofluorescent studies,a population of MSCs was obtained from human bone marrow aspiratesaccording to the protocol of Rickard et al. (Rickard D. J., et al.,1996, J. Bone Min. Res. 11: 312-324) and maintained in Control medium.To prevent spontaneous differentiation, cells were maintained atsubconfluent levels.

Indirect Immunofluorescence of Stem Cells

[0173] Stem cells and MSCs obtained from human bone marrow aspirateswere plated onto glass chamber slides and fixed for 15 minutes in 4%paraformaldehyde in 100 mM sodium phosphate buffer (pH 7.0). The cellswere washed for 10 minutes in 100 mM glycine in PBS (PBS/glycine) andblocked for 1 hour in Immunofluorescent Blocking Buffer (IBB; 5% BSA,10% FBS, 1×PBS, 0.1% Triton X-100). The cells were subsequentlyincubated for 1 hour in IBB containing the following cell-specificmonoclonal antibodies: 1) anti-smooth muscle actin (anti-SMA; CedarlaneInc, Homby Ont), to identify smooth muscle cells and pericytes (Skalli,O., et al., 1986, J. Cell Biol. 103:2787-2796; Schurch, W., et al.,1987, Am J. Pathol 128:91-103; Nehls, A. and D. Drenckhahn 1991, J.CellBiol. 113:147-154; Barghom, A. et al., 1998, Pediatr. Pathol. Lab.Med. 18:5-22)); 2) anti-Factor VIII (anti-FVIII; Calbiochem, San Diego,Calif.), to identify endothelial cells (Jaffe, E A, et al., 1973, J.Clin. Envest. 52:2757-2764; Nagle, R B, et al., 1987 Lymphology20:20-24); and 3) ASO2 (dianova, Hamburg, Germany), to identifyfibroblasts and cells of mesenchymal origin (Saalbach, A., et al., 1996J. Invest.Dermatol. 106:1314-1319; Saalbach, A., et al., 1997 Cell andTiss. Res. 290:595-599). The cells were washed extensively withPBS/glycine and incubated for 1 hour in IBB containing anFITC-conjugated secondary antibody. The cells were washed withPBS/glycine and mounted with a solution containing DAPI to visualizenuclei (VectaShield, Vector Labs, Burlingame, Calif.).

Flow Cytometry

[0174] Adipose-derived stem cells samples from 5 donors were cultured inControl Medium for 72 hours prior to analysis. Flow cytometry wasperformed on a FACScan argon laser cytometer (Becton Dickson, San Jose,Calif.). Cells were harvested in 0.25% trypsin/EDTA and fixed for 30minutes in ice-cold 2% formaldehyde. Following fixation, cells werewashed in Flow Cytometry Buffer (FCB; 1×PBS, 2% FBS, 0.2% Tween-20).Cell aliquots (1×10⁶ cells) were incubated in FCB containing monoclonalantibodies to Factor VIII, smooth muscle actin or ASO2. In addition,cells were also incubated with FCB containing a monoclonal antibody tovimentin (anti-VIM; Biogenesis, Brentwood, N.H.), to identifymesenchymal cells (Lazarides, E. 1982 Annu. Rev. Biochem. 51:219-250;Suza, S., et al., 1996 Eur. J. Cell Biol. 70:84-91). To assessviability, duplicate samples were harvested, fixed for 30 minutes withice-cold 1% paraformaldehyde, permeabilized with 0.05% Nonidet-40 andincubated with propidium iodide (PI) at a concentration of 25 μg/ml.Debris and dead cells were excluded by eliminating PI-positive events.All subsequent adipose-derived stem cell samples were correctedaccordingly.

Cumulative Population Doubling

[0175] Adipose-derived stem cells cells were maintained in ControlMedium until 80% confluent. Cells were harvested at confluence andpopulation doubling calculated using the formula log N₁/logN₂, where N₁is the number of cells at confluence prior to passaging and N₂ is thenumber of cells seeded after passaging. Cumulative population doublingwas determined in cultures maintained until passage 13 (approximately165 days). The mean cumulative population doubling obtained from 3donors was expressed as a function of passage number.

Cell Senescence Assay

[0176] Senescence was assessed using a β-gal staining assay, in whichβ-galactosidase activity is detected in senescent cells at pH 6.0 but isabsent in proliferating cells (Dimri, G P, et al., 1995 Proc. Natl.Acad. Sci. USA 92:9363-9367). Cells from each culture passage (passage 1to passage 15) were fixed for 5 minutes in 2%formaldehyde/glutaraldehyde and incubated in a β-Gal Reaction Buffer (1mg/ml X-Gal, 40 mM citric acid/sodium phosphate buffer (pH 6.0), 5 mMeach of potassium ferrocyanide and potassium ferricyanide, 150 mM NaCland 2 mM MgCl₂). Senescent cells (blue) were identified by lightmicroscopy.

Confirmation of Multi-lineage Differentiation of Adipose-Derived StemCells

[0177] Adipose-derived stem cells at passage 1 were analyzed for theircapacity to differentiate toward the adipogenic, osteogenic,chondrogenic and myogenic lineages. To induce differentiation, the stemcells were cultured with specific induction media as detailed inTable 1. Each media has been previously described and shown to inducemulti-lineage differentiation of MSCs (Pittenger, M F., et al., 1999Science 284:143-147; Grigoradis, A., et al., 1988 J. Cell Biol.106:2139-2151; Cheng, S-L., et al., 1994 Endo 134:277-286; Loffler, G.,et al., 1987 Klin. Wochenschr. 65:812-817; Hauner, H., et al., 1987 J.Clin. Endocrinol. Metabol. 64:832-835). Differentiation was confirmedusing the histological and immunohistological assays outlined in Table2. A commercial source of bone marrow-derived MSCs and lineage-specificprecursors were examined as positive controls. The adipose-derived stemcells were maintained in Control Medium and HFFs were analyzed asnegative controls.

[0178] 1. Adipogenesis: Adipogenic differentiation was induced byculturing the stem cells for 2 weeks in Adipogenic Medium (AM) andassessed using an Oil Red O stain as an indicator of intracellular lipidaccumulation (Preece, A. 1972 A Manual for Histologic Technicians,Boston, Mass.: Little, Brown, and Co.). Prior to staining, the cellswere fixed for 60 minutes at room temperature in 4% formaldehyde/1%calcium and washed with 70% ethanol. The cells were incubated in 2%(w/v) Oil Red O reagent for 5 minutes at room temperature. Excess stainwas removed by washing with 70% ethanol, followed by several changes ofdistilled water. The cells were counter-stained for 2 minutes withhematoxylin.

[0179] 2. Osteogenesis: Osteogenic differentiation was induced byculturing the stem cells for a minimum of 2 weeks in Osteogenic Medium(OM) and examined for Alkaline Phosphatase (AP) activity and ECMcalcification by von Kossa staining. To detect AP activity, cells wereincubated in OM for 2 weeks, rinsed with PBS and stained with a 1% APsolution (1% napthol ABSI phosphate, 1 mg/ml Fast Red TR) at 37° C. for30 minutes. For von Kossa staining, the cells were incubated in OM for 4weeks and fixed with 4% paraformaldehyde for 60 minutes at roomtemperature. The cells were rinsed with distilled water and thenoverlaid with a 1% (w/v) silver nitrate solution in the absence of lightfor 30 minutes. The cells were washed several times with distilled waterand developed under UV light for 60 minutes. Finally, the cells werecounter-stained with 0.1% eosin in ethanol.

[0180] 3. Chondrogenesis: Chondrogenic differentiation was induced usingthe micromass culture technique (Ahrens, P B, et al., 1977 Develop.Biol. 60:69-82; Reddi, A H 1982 Prog. Clin. Biol. Res. 110 (partB):261-268; Denker, A E., et al., 1995 Differentiation 59:25-34).Briefly, 10 μl of a concentrated adipose-derived stem cell suspension(8×10⁶ cells/ml) was plated into the center of each well and allowed toattach at 37° C. for two hours. Chondrogenic medium (CM) was gentlyoverlaid so as not to detach the cell nodules and cultures weremaintained in CM for 2 weeks prior to analysis. Chondrogenesis wasconfirmed using the histologic stain Alcian Blue at acidic pH. The stemcell nodules were fixed with 4% paraformaldehyde for 15 minutes at roomtemperature and washed with several changes of PBS. Studies have shownspecific staining of sulfated proteoglycans, present in cartilagenousmatrices, at pH levels of 1 and below (Lev, R. and S. Spicer 1964 J.Histochem. Cytochem. 12:309-312). In light of this, the cells wereincubated for 30 minutes with 1% (w/v) Alcian Blue (Sigma A-3157) in0.1N HCl (pH 1.0) and washed with 0.1N HCl for 5 minutes to removeexcess stain. In addition to Alcian Blue staining, expression of thecartilage-specific collagen type II isoform was also determined. Thestem cells were fixed in 4% paraformaldehyde for 15 minutes at roomtemperature. Cells were incubated in 0.2 U/ml chondroitinase ABC for 40min at 37° C. to facilitate antibody access to collagen II. The cellswere rinsed in PBS and endogenous peroxidase activity quenched byincubating for 10 minutes in 3% hydrogen peroxide in methanol. Followinga wash in PBS, non-specific sites were blocked by incubating cells for 1hour in Blocking Buffer (PBS, containing 10% Horse Serum). The cellswere subsequently incubated for 1 hour in Blocking Buffer containing amonoclonal antibody specific to human collagen type II (ICN Biomedical,Costa Mesa, Calif.). The cells were washed extensively in BlockingBuffer and collagen type II visualized using a commercially availablekit for the detection of monoclonal antibodies according to themanufacturer (VectaStain ABC kit, Vector Labs Inc., Burlingame, Calif.).

[0181] 4. Myogenesis: Myogenic differentiation was induced by culturingthe adipose-derived stem cells in Myogenic Medium (MM) for 6 weeks andconfirmed by immunohistochemical staining for the muscle-specifictranscription factor, MyoD1 and the myosin heavy chain. Cells wererinsed twice with PBS, fixed for 20 minutes with 4% paraformaldehyde andwashed several times with PBS. The cells were incubated with 3% hydrogenperoxide in PBS for 10 minutes to quench endogenous peroxidase enzymeactivity and non-specific sites were blocked by incubation in BlockingBuffer (PBS, 10% HS, 0.1% Triton X-100) for an additional 60 minutes.The cells were washed 3 times for 5 minutes each in Blocking Buffer andincubated for 1 hour in Blocking Buffer containing a either a monoclonalantibody specific to skeletal muscle myosin heavy chain (Biomeda, FosterCity, Calif.) or to MyoD1 (Dako Corp, Carpenteria, Calif.). The cellswere washed extensively in Blocking Buffer and the monoclonal antibodiesvisualized using the VectaStain ABC kit according to manufacturer'sspecifications. The cells were counter-stained with hematoxylin for 3minutes.

Results

[0182] Human adipose tissue was obtained by suction-assisted lipectomy(i.e. liposuction) and the lipoaspirates were processed based on adaptedmethodologies (Katz, A J, et al., 1999 Clin. Plast. Surg. 26:587-603,viii), in order to obtain a Processed Lipoaspirate or PLA cell(adipose-derived stem cells) population, containing the putative stemcell fraction. Processing of 300 cc of liposuctioned tissue routinelyyielded stem cell samples of 2-6×10⁸ cells. The cultures were maintainedin DMEM supplemented with 10% Fetal Bovine Serum (FBS). Supplementationwith FBS has been shown to be important for human and animal MSCattachment and proliferation in vitro (Haynesworth, S E, et al., 1992Bone 13:81-88; Lennon, D P, et al., 1995 Exp. Cell Res. 219:211-222;Lennon, D P, et al., 1996 In Vitro Cell Dev. Biol. 32:602-611). However,studies suggest that proliferation and differentiation of human MSCs maybe dependent upon FBS source and quality, making sera screening critical(Lennon, D P, et al., 1995 Exp. Cell Res. 219:211-222; Lennon, D P, etal., 1996 In Vitro Cell Dev. Biol. 32:602-611). The stem cells expandedeasily in vitro and exhibited a fibroblast-like morphology, consistentwith that of MSCs obtained from bone marrow and a commercial source(FIG. 1A). The stem cells did not appear to require specific sera lotsfor expansion and multi-lineage differentiation. Ten FBS lots from threemanufacturers were tested and did not appear to alter the stem cellmorphology, proliferation rate or their differentiative capacity invitro.

Growth Kinetics and Composition of the PLA

[0183] The adipose-derived stem cells, obtained from 20 donors andcultured under standard conditions (i.e. 10% FBS), exhibited an averagepopulation doubling time of 60 hours using several sera sources andlots. Following isolation, an initial lag time of 5 to 7 days wasobserved in stem cell cultures. Cells then entered a proliferative phasereaching confluence within 48 hours. To examine long-term growthkinetics of the stem cell cultures, we measured cumulative populationdoublings with respect to passage number in multiple donors. Consistentwith the observed lag time upon initial culture, the stem cellsunderwent an average of three population doublings prior to the firstpassage (FIG. 1B). An average of 1.5 population doublings was observedupon subsequent passages. A linear relationship between cumulativepopulation doubling and passage number was observed, indicating arelatively constant population doubling rate over the range studied.Furthermore, no appreciable decrease in cumulative population doublingswas observed at later passages (P13=165 days in culture), suggestingthat the stem cell cultures maintain their proliferative potentialduring extended culture periods.

[0184] In addition to cumulative population doubling, we also examinedcell senescence in long-term stem cell cultures using a β-gal stainingprotocol, in which β-galactosidase expression is absent in proliferatingcells but can be detected in senescent cells at a pH of 6.0 (Dimri, G P,et al., 1995 Proc. Natl. Acad. Sci. USA 92:9363-9367). Using this assay,the stem cell cultures were examined for senescence at each passage. Thestem cell cultures at passage 1 exhibited no appreciable β-gal staining(FIG. 1C, P1). An increase in β-gal staining was observed at laterpassages, however the percentage of senescent cells remained below 5%through 10 passages and increased to 15% at passage 15. Taken together,the data indicates that adipose-derived stem cell samples are relativelystable over long-term culture, maintaining a consistent populationdoubling rate and exhibiting low levels of senescence.

[0185] The SVF processed from excised adipose tissue is a heterogenouspopulation including mast cells, endothelial cells, pericytes,fibroblasts and lineage-committed progenitor cells, or pre-adipocytes(Pettersson, P. et al., 1984 Acta Med. Scand. 215:447-453; Hauner, H.,et al., 1987 J. Clin. Endocrinol. Metabol. 64:832-835). These componentsmay also be present, together with the putative stem cell fractionobtained from liposuctioned adipose tissue. However, no literatureregarding this has been published. To phenotypically characterize thestem cells, samples from several donors were examined by indirectimmunofluorescence using antibodies specific to established cell-surfacemarkers. A bone marrow stromal fraction obtained from human marrowaspirates was also examined as a control. To identify endothelial cells,the stem cells were incubated with a monoclonal antibody to Factor VIII(Jaffe, E A, et al., 1973 J. Clin. Invest. 52:2757-2764; Nagle, R B, etal., 1987 Lymphology 20:20-24). Smooth muscle cells were identifiedusing a monoclonal antibody to smooth muscle actin (Lazarides, E. 1982Annu. Rev. Biochem. 51:219-250; Suza, S., et al., 1996 Eur. J. CellBiol. 70:84-91). This antibody has also been shown to react withtransitional pericytes (i.e. pericytes of pre- and post-capillaries) andthe contractile apparatus of pericytes committed to the smooth musclelineage (Nehls, A. and D. Drenckhahn 1991 J. Cell Biol. 113:147-154;Herman, I M and P A D'Amore 1985 J. Cell Biol. 101:43-52). Low levels ofendothelial cells, smooth muscle cells and pericytes were observed inthe stem cell fraction (FIG. 2). In comparison, no staining for thesemarkers was observed in processed bone marrow stromal samples. Inaddition to Factor VIII and smooth muscle actin, cells were alsoincubated with a monoclonal antibody (ASO2) specific to fibroblasts andmesenchymal cells (Saalbach, A., et al., 1996 J. Invest. Dermatol.106:1314-1319; Saalbach, A., et al., 1997 Cell and Tiss. Res.290:595-599). The majority of the stem cells and bone marrow stromalcells stained positively with ASO2, suggesting a mesenchymal origin(FIG. 2, ASO2 panels).

[0186] To quantitatively determine the stem cell composition, sampleswere analyzed by flow cytometry using the cell surface markers describedabove. The samples were obtained and cultured for 72 hours in ControlMedium. Cell size and granularity were measured using forward and sidescatter settings (FIG. 3A). The majority of the stem cell sample wascomprised of small, agranular cells. In addition, the stem cells wereincubated with monoclonal antibodies to Factor VIII, smooth muscle actinand ASO2 and a monoclonal antibody to vimentin, an intermediate filamentprotein found predominantly in cells of mesenchymal origin (Lazarides,E. 1982 Annu. Rev. Biochem. 51:219-250; Suza, S., et al., 1996 Eur. J.Cell Biol. 70:84-91). Viability was assessed using propidium iodide andsamples were corrected for viability, non-specific fluorescence andautofluorescence. Data from a representative patient is shown (FIG. 3B).Cytometry data was collected from 5 donors and the number of positiveevents for each cell-specific marker was expressed as a percentage ofthe total stem cell number. Consistent with the immunofluorescent data,a fraction of the stem cells expressed Factor VIII (FVIII-positivecells=24.9%±8.2 of total stem cell number) and smooth muscle actin(SMA-positive cells=29.2%±2.1 of total PLA cell number) (FIG. 3C),indicating that the stem cell fraction contains endothelial cells,smooth muscle cells and, possibly, pericytes. Furthermore, the majorityof the stem cells stained positively for ASO2 (ASO2-positivecells=85.0%±12.8 of total PLA cell number) and vimentin (VIM-positivecells=63.2%±5.6 of total cell number), indicative of cells ofmesenchymal origin. Taken together, the results suggest that the stemcell fraction is a relatively homogenous population of mesodermal ormesenchymal cells with low contamination by endothelial cells, pericytesand smooth muscle cells.

Adipose-Derived Stem Cells Exhibit Multi-Lineage Potential

[0187] To study the multi-lineage capacity of the adipose-derived stemcells cells, cells were differentiated toward the adipogenic,osteogenic, chondrogenic and myogenic lineages using lineage-specificinduction factors (Table 1). Human and animal bone marrow-derived MSCshave been shown to differentiate toward the adipogenic, osteogenic andchondrogenic lineages with appropriate medium supplementation(Pittenger, M F., et al., 1999 Science 284:143-147; Grigoradis, A., etal., 1988 J. Cell Biol. 106:2139-2151; Cheng, S-L., et al., 1994 Endo134:277-286; Loffler, G., et al., 1987 Klin. Wochenschr. 65:812-817;Hauner, H., et al., 1987 J. Clin. Endocrinol. Metabol. 64:832-835).Following induction, differentiation was assessed using histology andimmunohistochemistry (Table 2). Commercially available MSCs andlineage-committed progenitor cells served as positive controls while thestem cells maintained in Control Medium and HFF cells were examined asnegative controls.

[0188] Pre-adipocytes and MSCs treated with adipogenic induction medium,containing cAMP agonists and induction agents such asisobutyl-methylxanthine (IBMX), indomethacin, insulin and dexamethasone,develop lipid-containing droplets that accumulate the lipid dye OilRed-O (Pittenger, M F., et al., 1999 Science 284:143-147; Rubin, C S, etal., 1978 J. Biol. Chem. 253:7570-7578; Deslex, S, et al., 1987 Int. J.Obesity 11:19-27). To determine if PLA cells undergo adipogenesis, cellswere cultured in medium containing these agents (Adipogenic Medium, AM)and stained with Oil Red-O. The stem cells cultured in AM werereproducibly induced toward the adipogenic lineage as early as two weekspost-induction (FIG. 4). A significant fraction of the cells containedmultiple, intracellular lipid-filled droplets that accumulated OilRed-O. The Oil Red O-containing stem cells exhibited an expandedmorphology with the majority of the intracellular volume (90-98%)occupied by lipid droplets, consistent with the phenotype of matureadipocytes. The mean level of adipogenic differentiation measured in 6donors under 35 years of age was 42.4%±10.6% (% Oil Red 0-positive cells/total PLA cell number). Prolonged culture times (i.e. 4 weeks) resultedin the detachment of differentiating cells from the culture plate aridflotation to the surface. The observed morphology and lipid accumulationof differentiated stem cells were similar to that observed upontreatment of bone marrow-derived MSCs and the pre-adipocyte cell line,3T3-L1, in AM. No lipid droplets were observed in undifferentiated stemcells or in HFF negative controls. In contrast to MSCs, in whichadipogenic differentiation dramatically decreases beyond the thirdculture passage (Conget, P A and J J Minguell 1999 J. Cell. Physiol.181:67-73), the adipogenic potential of the stem cells was maintainedover long-term culture (i.e. passage 15=175 days culture). Takentogether, the results indicate that the stem cells undergo adipogenicdifferentiation.

[0189] Differentiation of osteoprogenitor cells and marrow-derived MSCsinto osteoblasts is induced in vitro by treating cells with lowconcentrations of ascorbic acid, β-glycerophosphate and dexamethasone(Pittenger, M F, et al., 1999 Science 284:143-147; Cheng, S-L, et al.,1994 Endo 134:277-286; Conget, P A and J J Minguell 1999 J. Cell.Physiol. 181:67-73). Early differentiation of these cells into immatureosteoblasts is characterized by Alkaline Phosphatase (AP) enzymeactivity with human MSCs expressing AP as early as 4 days and maximumlevels observed at 12 days post-induction (Jaiswal, N, et al., 1997 J.Cell Biochem. 64:295-312). To confirm their osteogenic capacity, thestem cells were treated with osteogenic medium (OM) for 14 days and theexpression of AP was examined. The stem cells cultured in OM formed anextensive network of dense, multi-layered nodules that stainedpositively for AP (FIG. 5). The mean number of AP-positive stainingcells measured in 6 donors was 50.2%±10.8% of total stem cell number.Expression of AP was also observed in both MSCs and NHOst positivecontrols maintained in OM. In contrast, undifferentiated stem cells andHFF negative controls did not show evidence of AP expression. While APexpression is dramatically upregulated in osteogenic tissues, itsexpression has been observed in several non-osteogenic cell types andtissues such as cartilage, liver and kidney (Henthom, P S, et al., 1988J. Biol. Chem. 263:12011; Weiss, M J, et al., 1988 J. Biol. Chem.263:12002; Leboy, P S, et al., 1989 J. Biol. Chem. 264:17281).Therefore, AP expression is frequently used, in conjunction with otherosteogenic specific markers, as an indicator of osteogenesis. One suchindicator is the formation of a calcified extracellular matrix (ECM).Mature osteoblasts secrete a collagen I-rich ECM that becomes calcifiedduring the later stages of differentiation (Scott, D M 1980 Arch.Biochem. Biophys. 201:384-391). Therefore, in order to confirmosteogenic differentiation, calcification of the ECM matrix was assessedin the stem cells using a von Kossa stain. Calcification appears asblack regions within the cell monolayer. Consistent with osteogenesis,several black regions, indicative of a calcified ECM, were observed inthe stem cells treated for 4 weeks in OM. Calcification was alsoidentified in MSC and NHOst positive controls, while no calcificationwas observed in undifferentiated stem cells or HFF cells. The osteogenicpotential of the stem cells was maintained over long-term culture, withcells expressing AP as late as 175 days of culture. Taken together, theexpression of AP by the adipose-derived stem cells and the production ofa calcified ECM strongly suggests that these adipose-derived cells canbe induced toward the osteogenic lineage.

[0190] Chondrogenic differentiation can be induced in vitro using amicromass culture technique, in which cellular condensation (a criticalfirst event of chondrogenesis) is duplicated (Ahrens P. B., et al., 1977Develop. Biol. 60: 69-82; Reddi A. H. 1982 Prog. Clin. Biol. Res. 110 PtB: 261-268; Denker, A. E., et al., 1995 Differentiation 59, 25-34;Tacchetti, C, et al., 1992 Exp Cell Res. 200:26-33). Enhanceddifferentiation can be obtained by treating cells with dexamethasone andTGFβ1 (Iwasaki, M. et al., 1993 Endocrinology 132:1603-1608).Marrow-derived MSCs, cultured with these agents under micromassconditions, form cell nodules associated with a well-organized ECM richin collagen II and sulfated proteoglycans (Pittenger, M F, et al., 1999Science 284:143-147; Mackay, A M, et al., 1998 Tissue Eng. 4:415-428).These sulfated proteoglycans can be specifically detected using thestain Alcian Blue under acidic conditions (Lev, R and S. Spicer 1964 J.Histochem. Cytochem. 12:309-312). To assess the chondrogenic capacity ofthe stem cells, the cells were cultured via micromass in ChondrogenicMedium (CM), containing dexamethasone and TGFβ1. Micromass culture ofthe stem cells resulted in the formation of dense nodules consistentwith chondrogenic differentiation. The stem cell nodules were associatedwith an Alcian Blue-positive ECM, indicative of the presence of sulfatedproteoglycans within the matrix (FIG. 6). Cartilaginous nodules werealso observed upon micromass culture of MSC controls. To confirm thespecificity of Alcian Blue for cartilaginous matrices, human cartilageand bone sections were stained with Alcian Blue under acidic conditions.As expected, human cartilage sections stained positively with AlcianBlue, while no staining was observed in bone sections. In addition tothe presence of sulfated proteoglycans within the ECM, both stem cellsand human cartilage sections expressed the cartilage-specific collagentype II isoform, while no staining was observed in undifferentiated stemcells. Consistent with adipogenic and osteogenic differentiation, thestem cells retained their chondrogenic differentiation potential afterextended culture periods (i.e. up to 175 days). The above resultssuggest that the adipose-derived stem cells possess the capacity todifferentiate toward the chondrogenic lineage.

[0191] Myogenesis is characterized by a period of myoblastproliferation, followed by the expression of muscle-specific proteinsand fusion to form multinucleated myotubules. Early myogenicdifferentiation is characterized by the expression of several myogenicregulatory factors including Myogenic Determination factor1 (MyoD1;(Davis, R. L., et al., 1987 Cell 51:987-1000; Weintraub, H., et al.,1991 Science 251:761-763; Dias, P., et al., 1994 Semin. Diagn. Pathol.11:3-14). Terminally differentiated myoblasts can be characterized bythe expression of myosin and the presence of multiple nuclei(Silberstein, L., et al., 1986 Cell 46:1075-1081). Proliferation andmyogenic differentiation of muscle precursors and bone marrow-derivedstem cells can be induced by dexamethasone and results in the expressionof muscle-specific proteins (Grigoradis, A, et al., 1988 J. Cell Biol.106:2139-2151; Ball, E H and B D Sanwal 1980 J. Cell. Physiol 102:27-36;Guerriero, V and J R Florini 1980 Endocrinology 106:1198-1202).Furthermore, addition of hydrocortisone is known to stimulate humanmyoblast proliferation, prior to their transition into differentiatedmyotubules (Zalin, R J 1987 Exp. Cell Res. 172:265-281). To examine ifthe stem cells undergo myogenesis, the cells were cultured for 6 weeksin the presence of dexamethasone and hydrocortisone, and incubated withantibodies specific to MyoD1 and myosin (heavy chain). Consistent withearly myogenic differentiation, treatment of the stem cells with MM for1 week induced the expression of MyoD1 (FIG. 7). The stem cells treatedfor longer time periods (6 weeks) stained positively for myosin. Inaddition to myosin expression, the presence of discrete ‘patches’ oflarge, elongated cells with multiple nuclei were also observed,suggesting that the stem cells underwent myoblast fusion (PLA panel,inset). MyoD1 and myosin heavy chain expression were also detected inhuman skeletal muscle positive control cells. Using Myogenic Medium,myogenic differentiation was not observed in MSC controls even at 6weeks of induction. These cells may be adversely affected byhydrocortisone and may require alternate conditions to inducedifferentiation. Myogenic differentiation levels in the stem cellsaveraged 12%. Multi-nucleation, myosin heavy chain and MyoD1 expressionwere not observed in undifferentiated stem cells nor in HFF negativecontrols. The presence of multi-nucleated cells and the expression ofboth MyoD1 and myosin heavy chain suggests that the adipose-derived stemcells have the capacity to undergo myogenic differentiation. TABLE 1Lineage-specific differentiation induced by media supplementation MediumMedia Serum Supplementation Control DMEM 10% FBS none Adipogenic DMEM10% FBS 0.5 mM isobutyl-methylxanthine (AM) (IBMX), 1 μM dexamethasone,10 μM insulin, 200 μM indomethacin, 1% antibiotic/ antimycoticOsteogenic DMEM 10% FBS 0.1 μM dexamethasone, 50 μM (OM)ascorbate-2-phosphate, 10 mM β-glycerophosphate, 1% anti-biotic/antimycotic Chondrogenic DMEM  1% FBS 6.25 μg/ml insulin, 10ng/ml (CM) TGFβ1, 50 nM ascorbate-2- phosphate, 1% antibiotic/anti-mycotic Myogenic DMEM 10% FBS, 0.1 μM dexamethasone, 50 μM (MM)  5% HShydrocortisone, 1% antibiotic/anti- mycotic

[0192] TABLE 2 Differentiation markers and assays of lineage-specificdifferentiation Lineage-specific Histologic/immunohisto- Lineagedeterminant chemical assay Adipogenic Lipid accumulation Oil Red O stainOsteogenic 1. Alkaline phosphatase 1. Alkaline Phosphatase activitystain 2. Calcified matrix 2. Von Kossa stain production Chondrogenic 1.Sulfated proteoglycan- 1. Alcian Blue (pH 1.0) rich matrix stain 2.Collagen II synthesis 2. Collagen II-specific monoclonal antibodyMyogenic 1. Multi-nucleation 1. Phase contrast 2. Skeletal musclemicroscopy myosin heavy 2. Myosin & MyoD1 chain & MyoD1 specificmonoclonal expression antibodies

Discussion

[0193] Conceptually, there are two general types of stem cells:Embryonic Stem Cells (ESCs) and autologous stem cells. Althoughtheoretically appealing because of their pluripotentiality, thepractical use of ESCs is limited due to potential problems of cellregulation and ethical considerations. In contrast, autologous stemcells, by their nature, are immunocompatible and have no ethical issuesrelated to their use. For the engineering of mesodermally derivedtissues, autologous stem cells obtained from bone marrow have provenexperimentally promising. Human bone marrow is derived from theembryonic mesoderm and is comprised of a population of HematopoeiticStem Cells (HSCs), supported by a mesenchymal stroma (Friedenstein A.J., et al., 1968 Transplantation 6: 230-47; Friedenstein A. J., et al.,1974 Transplantation 17: 331-40; Werts E. D., et al., 1980 Exp. Hematol.8: 423; Dexter T. M. 1982 J. Cell Physiol. 1: 87-94; Paul S. R., et al.,1991 Blood 77: 1723-33). While the proliferation and differentiation ofHSCs has been well documented, less is known about the stromalcomponent. The bone marrow stroma, in both animals and humans, isheterogenous in composition, containing several cell populations,including a stem cell population termed Mesenchymal Stem Cells or MSCs(Caplan A I 1991 J. Orthop. Res. 9:641-650). Studies on MSCs havedemonstrated their differentiation into adipocytes (Beresford J. N., etal., 1992 J. Cell Sci. 102; 341-351; Pittenger M. F., et al., 1999Science 284: 143-147), chondrocytes (Caplan A. I. 1991 J. Orthop. Res.9: 641-50; Pittenger M. F., et al., 1999 Science 284: 143-147; Berry L.,et al., 1992 J. Cell Sci. 101: 333-342; Johnstone B., et al., 1998 Exp.Cell Res. 238: 265-272; Yoo J. U., et al., 1998 J. Bone Joint Surg. Am.80: 1745-1757), myoblasts (Wakitani S., et al., 1995 Muscle Nerve 18:1417-1426; Ferrari G., et al., 1998 Science 279: 1528-1530) andosteoblasts (Caplan A. I. 1991 J. Orthop. Res. 9: 641-50; Pittenger M.F., et al., 1999 Science 284: 143-147; Grigoradis A., et al., 1988 J.Cell Biol. 106: 2139-2151; Cheng S-L., et al., 1994 Endo 134: 277-286,1994; Haynesworth S. E., et al., 1992 Bone 13: 81-8; Rickard D. J., etal., 1996 J. Bone Min. Res. 11: 312-324; Prockop D. J. 1997 Science 276:71-74; Dennis J. E., et al., 1999 J. Bone Miner. Res. 14: 700-709).These cells represent a promising option for future tissue engineeringstrategies. However, traditional bone marrow procurement procedures maybe painful, frequently requiring general or spinal anesthesia and mayyield low numbers of MSCs upon processing (approximately 1 MSC per 10⁵adherent stromal cells (Pittenger, M F et al., 1999 Science 284:143-147;Rickard, D J, et al., J. Bone Min. Res. 11:312-324: Bruder, S P, et al.,1997 J. Cell. Biochem. 64:278-294)). From a practical standpoint, lowstem cell numbers necessitate an ex vivo expansion step in order toobtain clinically significant cell numbers. Such a step is timeconsuming, expensive and risks cell contamination and loss. An idealsource of autologous stem cells would, therefore, be both easy toobtain, result in minimal patient discomfort yet be capable of yieldingcell numbers substantial enough to obviate extensive expansion inculture.

[0194] We report that a cellular fraction with multiple mesodermallineage capabilities can be processed from human lipoaspirates. Thiscellular fraction is the adipose-derived stem cells which is designateda Processed Lipoaspirate (PLA), comprising fibroblast-like cells thatcan be expanded easily in vitro without the need for specific sera lotsor media supplementation. The stem cell samples maintained a lineargrowth rate with no appreciable senescence over extended cultureperiods. The stem cell population was heterogenous in nature, with themajority of the cells being mesenchymal in origin. However,contaminating endothelial, smooth muscle and pericyte cell populationswere identified. The stem cells also exhibited multi-lineage potentialin vitro, differentiating toward the adipogenic, osteogenic,chondrogenic and myogenic lineages when cultured in the presence ofestablished lineage-specific differentiation factors. Thedifferentiation results were consistent with those observed uponlineage-specific differentiation of bone marrow-derived MSCs andlineage-committed precursors.

[0195] While the apparent multi-differentiative capacity of the stemcells suggests the presence of a stem cell population within humanliposuctioned adipose tissue, it is not conclusive. Multi-lineagedifferentiation may also be due to the presence of: (1) multiplelineage-committed progenitor cells; (2) multi-potent cells from othersources (e.g. pericytes, marrow-derived MSCs from peripheral blood); or(3) a combination of the above.

[0196] The observed differentiation may be due to the presence oflineage-committed progenitor cells, such as pre-osteoblasts,pre-myoblasts or pre-adipocytes within the stem cell fraction. Cellularfractions (i.e. SVFS) obtained from excised adipose tissue are known tocontain pre-adipocytes that differentiate into mature adipocytes(Pettersson, P, et al., 1984 Acta Med. Scand. 215:447-453; Pettersson,P, et al., 1985 Metabolism 34:808-812). It is possible that the observedadipogenic differentiation by the stem cells is simply the commitment ofexisting pre-adipocytes and not the differentiation of a multi-potentcell. However, we do not believe this to be the case. As little as 0.02%of the SVF obtained from excised adipose tissue have been identified aspre-adipocytes capable of adipogenic differentiation (Pettersson, P, etal., 1984 Acta Med. Scand. 215:447-453). If pre-adipocyte numbers withinthe stem cell fraction are comparable to those levels measured in theSVF from excised tissue, one would expect a relatively low level ofadipogenesis. However, the degree of adipogenesis observed in the stemcells is significant (approximately 40% of the total PLA cell number)and may result from the differentiation of additional cell types.

[0197] Damage to the underlying muscle during liposuction may introducemyogenic precursor cells or satellite cells into the stem cell fraction,resulting in the observed myogenic differentiation by these cells.Located between the sarcolemma and the external lamina of the musclefiber, myogenic precursor cells in their undifferentiated state arequiescent and exhibit no distinguishing features, making theiridentification difficult. Several groups have attempted to identifyunique markers for these precursors with limited success. Currently, theexpression of the myogenic regulatory factors, MyoD1 and myogenin havebeen used to identify satellite cells during embryogenesis and inregenerating adult muscle in rodents (Cusella-DeAngelis, M. C., et al.,1992 Cell Biol. 116:1243-1255; Grounds, M. D., et al., 1992 Cell Tiss.Res. 267:99-104; Sassoon, D. A. 1993 Develop. Biol. 156:11; Maley, M. A.L., et al., 1994 Exp. Cell Res. 211:99-107; Lawson-Smith, M. J. andMcGeachie, J. K. 1998 J. Anat. 192:161-171). In addition, MyoD1expression has been identified in proliferating myoblasts prior to theonset of differentiation (Weintraub, H, et al., Science 251:761-763).While these markers have not been used to identify myogenic precursorsin human subjects, MyoD1 is expressed during early myogenicdifferentiation in normal skeletal muscle and has been used to identifythe skeletal muscle origin of rhabdosarcomas in humans 77-79 (Dias, P.,et al., 1990 Am. J. Pathol. 137:1283-1291; Rosai, J., et al., 1991 Am.J. Surg. Pathol. 15:974; Nakano, H., et al., 2000 Oncology 58:319-323).The absence of MyoD1 expression in the stem cells maintained innon-inductive Control Medium (see FIG. 28), suggests that our observedmyogenic differentiation is not due to the presence of myogenicprecursors or proliferating myoblasts within the stem cell fraction.Consistent with this, the blunt contour of the liposuction cannula wouldmake it extremely difficult to penetrate the fibrous fascial cavitysurrounding the muscle and introduce these precursors into the adiposecompartment.

[0198] Human adipose tissue is vascularized and, as such, containspotential systemic vascular ‘conduits’ for contamination by multi-potentcells, such as pericytes and marrow-derived MSCs. Disruption of theblood supply during liposuction may result in the release of pericytes,known to possess multi-lineage potential both in vivo and in vitro(Schor, A M, et al., 1990 J. Cell Sci. 97:449-461; Doherty, M J 1998 J.Bone Miner. Res. 13:828-838; Diefenderfer, D L and C T Brighton 2000Biochem. Biophys. Res. Commun. 269:172-178). Consistent with this, ourimmunofluorescent and flow cytometry data show that a small fraction ofthe stem cells is comprised of cells that express smooth muscle actin, acomponent of transitional pericytes and pericytes committed to thesmooth muscle lineage (Nehls, A. and D Drenckhahn 1991 J. Cell Biol.113:147-154). The multi-lineage differentiation observed in the stemcells may be, in part, due to the presence of pericytes. Disruption ofthe blood supply may also introduce MSCs into the stem cell fraction.However, the literature is conflicted as to the presence of these stemcells in the peripheral blood Huss, R 2000 Stem Cells 18:1-9; Lazaras, HM, et al., 1997 J. Hematother. 6:447-455). If the peripheral blood doesindeed represent a source of MSCs, our observed multi-lineagedifferentiation may be due to the contamination of adipose tissue bythese stem cells (MSCs). However, MSCs are a small constituent of thebone marrow stroma in humans (approximately 1 MSC per 10⁵ adherentstromal cells (Pittenger, M F, et al., 1999 Science 284:143-147;Rickard, D J 1996 J. Bone Min. Res. 11:312-324; Bruder, S P, et al.,1997 J. Cell. Biochem. 64:278-294). If these cells do exist inperipheral blood, they are likely to be in even smaller quantities thanin the bone marrow and contamination levels of the adipose-derived stemcells fraction by these cells may be negligible.

[0199] While these arguments may provide support for the presence of amulti-potent stem cell population within liposuctioned adipose tissue,definitive confirmation requires the isolation and characterization ofmultiple clones derived from a single cell. Preliminary data confirmsthat clonal stem cell populations possess multi-lineage potential,capable of adipogenic, osteogenic, and chondrogenic differentiation.

[0200] Current research has demonstrated positive results using bonemarrow-derived MSCs. MSCs can differentiate into osteogenic andchondrogenic tissues in vivo (Benayahu, D. et al., 1989 J. Cell Physiol140:1-7; Ohgushi, H M 1990 Acta Orthop. Scand. 61:431-434; Krebsbach, PH, et al., 1997 Transplantation 63:1059-1069; Bruder, S P, et al., 1998J. Orthop. Res. 16:155-162) and preliminary data suggests that thesecells can be used to repair bony and cartilagenous defects (Wakitani,S., et al., 1995 Muscle Nerve 18:1417-1426; Krebsbach, P H, et al., 1997Transplantation 63:1059-1069; Bruder, S P, et al., 1998 J. Orthop. Res.16:155-162; Bruder, et al., 1998 Clin. Orthop. S247-56; Krebsbach, P H1998 Transplantation 66:1272-1278; Johnstone and Yoo 1999 Clin. Orthop.S156-162). The stem cells obtained from liposuctioned adipose tissue mayrepresent another source of multi-lineage mesodermal stem cells. Likethe bone marrow stroma, these data suggests that adipose tissue maycontain a significant fraction of cells with multi-lineage capacity.These adipose-derived stem cells may be readily available in largequantities with minimal morbidity and discomfort associated with theirharvest.

EXAMPLE 8

[0201] The following description provides adipose-derived stem cellswhich differentiate into osteogenic tissue, and methods for isolatingsaid stem cells. The osteogenic potential of the stem cells decreaseswith the age of the donor. However, adipogenesis is not affected by ageof the donor.

Materials and Methods Lipoaspirate Collection and Processing

[0202] Human adipose tissue was obtained from elective liposuctionprocedures under local anesthesia according to patient consent protocolHSPC #98-08 011-02 (University of California Los Angeles). The rawlipoaspirate was processed to obtain the adipose-derived stem cellspopulation (Zuk, P, et al., 2001 Tissue Engineering 7:209-226). Briefly,raw lipoaspirates were washed extensively with equal volumes ofPhospho-Buffered Saline (PBS) and the extracellular matrix (ECM) wasdigested at 37° C. for 30 minutes with 0.075% collagenase. Enzymeactivity was neutralized with Dulbecco's Modified Eagle's Medium (DMEM;Life Technologies), containing 10% Fetal Bovine Serum (FBS; HyClone) andcentrifuged at 1200×g for 10 minutes. The stem cell pellet wasresuspended in DMEM/10% FBS and filtered through a cell strainer toremove any remaining tissue. The cells were incubated overnight at 37°C., 5% CO₂ in non-inductive control medium (DMEM, 10% FBS, 1%antibiotic/antimycotic solution). Following incubation, the plates werewashed extensively with PBS to remove residual non-adherent red bloodcells. The stem cells were maintained at 37° C., 5% CO₂ in controlmedium (Table 3). To prevent spontaneous differentiation, cultures weremaintained at sub-confluent levels. TABLE 3 Lineage-SpecificDifferentiation Induced By Media Supplementation Medium Media SerumSupplementation Control DMEM 10% FBS none Adipogenic DMEM 10% FBS 0.5 mMisobutyl-methylxanthine (IBMX), 1 μM dexamethasone, 10 μM insulin, 200μM indomethacin, 1% antibiotic/antimycotic Osteogenic DMEM 10% FBS 0.1μM dexamethasone, 50 μM ascorbate-2-phosphate, 10 mM β-glycerophosphate,1% antibiotic/ antimycotic

Induction and Analysis of Differentiation

[0203] 1. Adipogenic Differentiation: PLA cells (passage 1) were seededinto six well plates (Costar, Cambridge, Mass.) at a density of 4×10⁴cells per well and cultured in control medium for 72 hours. To induceadipogenic differentiation, PLA cells were cultured for 2 weeks inAdipogenic Medium (Table 3). PLA cells, at the same density, weremaintained in control medium as a negative control.

[0204] Oil Red O Staining: Adipogenesis was confirmed at two weekspost-induction by staining with Oil Red O to identify intracellularlipid vacuoles. Cells were fixed for 60 minutes at room temperature in4% formaldehyde/1% calcium and washed with 70% ethanol. The cells wereincubated in 2% (w/v) Oil Red O reagent (Sigma, St Louis, Mo.) for 5minutes at room temperature. Excess stain was removed by washing with70% ethanol, followed by several changes of distilled water. The cellswere counter-stained for 1 minute with hematoxylin.

[0205] 2. Osteogenic Differentiation: PLA cells (passage 1) were seededinto six well plates at a density of 1×10⁴ cells per well and culturedfor 72 hours in control medium. Based on previous studies on bonemarrow-derived Mesenchymal stem cells (Pittenger, M F 1999 Science284:143-147), PLA cells were maintained for a minimum of two weeks inOsteogenic Medium (Table 3) to induce osteogenesis. PLA cells weremaintained in control medium as a negative control.

[0206] Alkaline Phosphatase Staining: Alkaline phosphatase (AP) activitywas examined at 14 days post-induction. PLA cells were rinsed with PBSand incubated for 30 minutes at 37° C. in 0.05M Tris-HCl (pH 9)containing 1% (v/v) of a 50 mg/ml solution of naphthol AS-Biphosphate(Sigma) dissolved in dimethyl sulfoxide (DMSO) and 1 mg/ml Fast Red TRsalt (Sigma). Following incubation, cells were fixed for 10 minutes withan equal volume of 8% paraformaldehyde, followed by a rinse withdistilled water.

[0207] Von Kossa Staining: Extracellular matrix calcification wasdetected at four weeks post-induction by von Kossa staining. PLA cellswere fixed with 4% paraformaldehyde at room temperature for 1 hour,followed by a 30 minute incubation with a 5% (w/v) silver nitratesolution (Sigma) in the absence of light. The cells were washed severaltimes with distilled water, developed under UV light for 60 minutes andcounter-stained with 0.1% cosin in ethanol. Matrix calcification wasidentified by the presence of black extracellular deposits.

Quantitation of Differentiation

[0208] Adipogenic and osteogenic differentiation levels in each donorwere quantified using a Zeiss Axioscope 2 microscope fitted with a Spot2 digital camera and a 2× objective (magnification 200×). The totalnumber of Oil Red O- and AP-positive cells (adipogenesis andosteogenesis, respectively) in duplicate samples from each donor werecounted within three consistent regions from each well (e.g., atpositions 3, 6 and 9 o'clock). The total number of positive-stainingcells was expressed as a percentage of total PLA cell number countedwithin each region. Values from the three regions were averaged to givethe mean differentiation level for each donor. The mean level ofdifferentiation was expressed with respect to patient age. Von Kossaidentifies regions of matrix production rather than individualdifferentiated cells, therefore this staining procedure was used toconfirm osteogenic differentiation.

Quantitation of Osteogenic Precursors within PLA Samples

[0209] In order to estimate the number of osteogenic precursors withinthe PLA population, cells with osteogenic activity were counted andrelated to patient age. Specifically, two age groups were examined:Group A=20 to 39 years and Group B=40 to 60 years. First-passage PLAcells (P1) were seeded onto 100 mm dishes, induced toward the osteogeniclineage as described above and stained for AP activity to confirmdifferentiation. Precursor number within each PLA sample was determinedby counting the number of AP-positive colony forming units (CFU/AP⁺).Based on a previous study, a minimum of ten AP-positive cells was usedto identify a CFU/AP⁺ (Long, M, et al., 1999 J. Gerontol. A. Biol. Sci.Med. Soc. 54:B54-62). The average number of CFU/AP⁺ was determined andexpressed with respect to age group. The optimal number of PLA cellsrequired for osteogenic differentiation was determined empirically(1×10⁴, 5×10⁴, 1×10⁵ and 5×10⁵ cells plated per dish). While osteogenicdifferentiation levels were greatest at 5×10⁵ cells per dish, confluencylevels prevented accurate colony counting. Data was therefore obtainedusing a density of 1×10⁵ cells per dish.

Growth Kinetics

[0210] To measure PLA cell growth kinetics (population doubling) withrespect to donor age, PLA cells from each donor (P1) were seeded at adensity of 1×10⁴ cells into multiple dishes. Cell number was determinedfrom triplicate samples 24 hours after plating and every 48 hours untilday 11. A growth curve (cell number vs. culture time) was derived andpopulation doubling was calculated from the log phase.

Statistical Analysis

[0211] Significant differences in PLA cell osteogenesis and adipogenesisaccording to donor age were determined by linear regression analysis (rvalue). In addition, the mean levels of differentiation across donor agewere compared using an unpaired student t-test (assuming unequalvariances) and a one way analysis of variance (ANOVA).

Results PLA Cell Growth Kinetics Vary with Respect to Donor Age

[0212] Initial PLA cultures were relatively homogenous in appearance,with the majority of cells (85 to 90%) exhibiting a fibroblast-likemorphology. A small fraction of endothelial cells, macrophages andpre-adipocytes could be identified (less than 10% of the totalpopulation). PLA cells reached 80-90% confluency within 14 days ofculture in control medium. Growth curves derived from first-passage PLAcell cultures (P1) were characterized in each donor by an initial lagphase (typically 48 hrs), followed by a log phase (average=7 days) and aplateau phase. Representative growth kinetic curves are shown in FIG.8A. No significant difference in the duration of the lag and log phaseswas observed in any donor. Similarly, no significant differences in PLAgrowth kinetics were observed in younger patients (20 years vs. 39years). However, a decrease in the log phase of PLA cells was observedin older patients (eg. day 13; 58 years—12.6×10⁴ cells, 20years—26.9×10⁴ cells). Based on the growth kinetics data, the averagePLA cell population doubling time calculated from 15 donors was52.67±8.67 hours. PLA cell population doubling time ranged from 38 to 77hours (FIG. 8B; 20 years vs. 53 years). Regression analysis ofpopulation doubling and donor age yielded a positive correlation ofr=0.62 (n=15), indicating a trend toward increasing population doubling(i.e. decreasing proliferative potential) with age. However, statisticalanalysis of donors grouped according to decade (i.e. 20-30 years, 30-40years, 40-50 years, 50-60 years), using an unpaired t-test, did not showa significant difference in population doubling (p>0.05), suggestingthat PLA proliferation does not significantly decrease with increasingdonor age.

Adipogenic Differentiation Potential Does Not Change with PLA Age

[0213] Adipogenesis and lipid vacuole formation in PLA cells wereconfirmed by staining cells with the lipid dye Oil Red O. In all donors,low levels of adipogenic differentiation in PLA cells were apparent asearly as 5 days induction in Adipogenic Medium. Differentiating cellsassumed an expanded morphology consistent with adipocytes andaccumulated lipid-rich intracellular vacuoles that stained with Oil RedO (FIG. 9A, Panels 1 and 2). By 14 days post-induction, differentiatingcells contained large, Oil Red O-positive lipid droplets within thecytoplasm. Differentiation levels varied from donor to donor withseveral donors exhibiting low levels of adipogenesis, in whichindividual Oil Red O-positive cells containing a moderate amount ofstain were easily identified (FIG. 9A, Panel 1). In addition, severaldonors exhibited enhanced levels of adipogenesis, in which both thenumber of Oil Red O-positive cells and the accumulation of the stainincreased dramatically (FIG. 9A, Panel 2). Cells cultured innon-inductive control media exhibited no change in morphology and didnot accumulate Oil Red O, confirming the specificity of the inductivemedium conditions (FIGS. 9A, Panel 3). To measure changes in adipogenicdifferentiation potential with respect to donor age, the number of OilRed O-positive cells was directly counted within a defined region andexpressed as a percentage of the total number of PLA cells counted.Values from each region were averaged to give the mean level ofadipogenic differentiation and expressed with respect to donor age (FIG.9B). Adipogenic differentiation levels ranged from 4.51% to 57.78% ofthe total PLA cells (n=20). An average differentiation potential of26.55±18.14% was calculated. However, a negligible regression value wasobtained upon analysis (r=0.016), suggesting that no significant changesin adipogenic differentiation occur with increasing donor age.

Osteogenic Differentiation Decreases with Donor Age

[0214] To confirm osteogenesis, cells were stained for AlkalinePhosphatase (AP) activity and extracellular matrix calcification using asilver nitrate/Von Kossa stain. PLA cells, cultured in OsteogenicMedium, underwent a dramatic change in cellular morphology as early as 4days post-induction, changing from spindle-shaped to cuboidal,characteristic of osteoblasts. Low levels of osteogenesis werecharacterized in some donors by the formation of a monolayer ofAP-positive cells (FIG. 10A, Panel 1). Higher levels of osteogenesiswere characterized in some patients by the presence of multi-layeredAP-positive nodular structures with well-defined inter-nodular regionscontaining no cells (FIG. 10A, Panel 2). In addition to AP activity,regions of mineralization, as detected by von Kossa staining, wereevident after 3 weeks of culture, further substantiating osteogenicdifferentiation (FIG. 10A, Panels 4 and 5). Control PLA cells did notexhibit AP activity or matrix mineralization (FIG. 10A, Panels 3 and 6).To measure potential changes in osteogenic differentiation with donorage, the mean level of osteogenesis (i.e. AP-positive cells) wasdetermined using the same method to calculate adipogenic levels.

[0215] In contrast to adipogenesis, a significant decrease inosteogenesis was observed in older donors. Osteogenic differentiationranged from 11.64% to 64.69% of the total PLA cells (FIG. 10B).Regression analysis of donor age and osteogenesis yielded a significantnegative correlation (r=−0.70, n=19), suggesting that osteogenicdifferentiation decreases with respect to age. A similar trend wasobserved using von Kossa staining. Interestingly, a distinct decrease inosteogenic differentiation was observed in donors older than 36 years ofage (FIG. 10B, dashed line). Consistent with this, a significantdifference in osteogenesis (p<0.001) was observed when the subjects weredivided into two age groups. Donors from the younger age group (20 to 36years; n=7) exhibited a mean osteogenic potential of 50.7±10% (total PLAcells) while a significantly lower level of osteogenesis (20.7±7.9%total PLA cells) was measured in the older age group (37 to 58 years;n=11) (FIG. 10C). Based on this data, cells from the younger groupexhibited a 2.4-fold increase in osteogenic potential, forming 59% moreAP-positive cells.

Relative Proportion of Osteogenic Precursors Within PLA

[0216] In order to determine if the decrease in PLA osteogenesis is dueto a decrease in the number of PLA cells with osteogenic potential, therelative proportion of osteogenic precursor cells within the PLA wascalculated with respect to donor age. PLA cells were induced for 2 weeksin Osteogenic Medium and the number of precursors within the PLAdetermined by calculating the number of AP-positive Colony Forming Units(CFU/AP⁺) (Grigoradis, A, et al., 1988 J. Cell Biol. 106:2139-2151;Pittenger, M F, et al., 1999 Science 284:143-147; Jaiswal, N., et al.,1997 J. Cell Biochem. 64:295-312). The number of precursors wascalculated in two age groups (Group A=20-39 years, n=5 and Group B=40-58years, n=6). Consistent with the diminished osteogenic potentialobserved in older PLA samples, a slight decrease in CFU/AP⁺ number wasobserved with increasing age. The average number of CFU/AP⁺ in Group Awas 194±61 per 10⁵ PLA cells, while the number of CFU/AP⁺ in Group Bdecreased to 136±32 per 10⁵ PLA cells (FIG. 11). While a decreasingtrend in osteoprogenitor cells was observed, this decrease was notstatistically significant (p=0.11), suggesting that the decrease inosteogenic potential by PLA cells may not be directly due to a decreasein the number of osteogenic precursors.

Discussion

[0217] Mesenchymal stem cells can be isolated from bone marrow.Mesenchymal stem cells are a component of the bone marrow stroma andpossess the capacity to differentiate into various mesodermal tissuesincluding fat, bone and cartilage (Grigoradis, A., et al., 1988 J. CellBiol. 106:2139-2151; Caplan, A. 1. 1991 J. Orthop. Res. 9:641-650;Beresford, J. N., et al., 1992 J. Cell Sci. 102:341-351; Berry, L., etal., 1992 J. Cell Sci. 101:333-342; Ferrari, G., et al., 1998 Science279:1528-1530; Johnstone, B., et al., 1998 Exp. Cell Res. 238:265-272;Pittenger, M. F., et al., 1999 Science 284:143-147). This multi-lineagepotential may be clinically useful for the repair of complexpost-traumatic and congenital defects. Indeed, several in vitro and invivo studies have suggested the clinical potential for these stem cells(Benayahu, D., et al., 1989 J. Cell Physiol. 140:1-7; Wakitani, S., etal., 1995 Muscle Nerve 18:1417-1426; Krebsbach, P. H., et al., 1997Transplantation 63:1059-1069; Bruder, S. P., et al., 1998 Clin. Orthop.(355 Suppl):S247-56; Johnstone, B., and Yoo, J. U. 1999 Clin. Orthop.(367 Suppl):S156-62). However, bone marrow procurement is painful,requires general anesthesia and yields low numbers of mesenchymal stemcells upon processing (Pittenger, M. F., et al., 1999 Science284:143-147; Rickard, D J, et al., 1996 J. Bone Miner. Res. 11:312-324;Bruder, S P, et al., 1997 J. Cell. Biochem. 64:278-294), thus requiringan ex vivo expansion step prior to clinical use. In light of thesefactors, an additional source of multi-lineage stem cells may bedesirable. We have identified a population of stem cells in thestromal-vascular fraction of liposuctioned human adipose tissue (Example7, supra). This cell population is designated a Processed Lipoaspirate(PLA), and appears to be similar to bone marrow-derived mesenchymal stemcells in many aspects. Like mesenchymal stem cells, PLA cells are stableover long-term culture, expand easily in vitro and possess multi-lineagepotential, differentiating into adipogenic, osteogenic, myogenic andchondrogenic cells.

[0218] PLA cells possess a fibroblast-like morphology, expand stably invitro, and proliferate with an average population doubling time of 53hours. Previous studies have shown that the size and number ofadipocytes within adipose tissue increases with age (Hauner, H. et al.,1987 J. Clin. Endocrinol. Metabol. 64:832-835) suggesting an overallincrease in adipogenesis in the adipose stores with advancing age. Incontrast to these studies, we do not observe a significant age-relatedchange in adipogenesis by PLA cells, suggesting that the adipogenicpotential of older PLA cells is unaffected by advancing age. Thedevelopment of adipose tissue requires the activity of several growthfactors and steroid hormones (Hatmer, H. et al., 1987 J. Clin.Endocrinol. Metabol. 64:832-835). Therefore, the adipogenic potential ofPLA cells may be influenced by the genetic background and/or hormonallevels within each donor. Proenza et al. has reported that adipogenesiscan be affected by alterations in the expression of several genes,including lipoprotein lipase, adrenoreceptor and uncoupling protein(Rickard, D J, et al., 1996 J. Bone Miner. Res. 11:312-324; Glowacki J.1995 Calcif. Tiss. Int. 56 (Suppl 1):S50-51). In addition, Chen et al.has shown that the expression of specific obesity-related genes inpre-adipocytes is related to the differentiation of these cells intomature adipocytes (Chen, X., et al., 1997 Biochim. Biophys.1359:136-142). Therefore, gene expression levels, together with hormonalactivity, may differ from donor to donor, influencing the adipogenicpotential of PLA cells and resulting in varying levels of adipogenesis,irrespective of donor age.

[0219] In contrast to adipogenesis, a decrease in PLA osteogenicpotential (as measured by AP activity) is observed with increasing donorage. A significant negative correlation between osteogenesis and donorage is found by regression analysis (r=−0.70). Furthermore, asignificant difference in osteogenesis is observed when donors aresegregated into two age groups (20 to 36 years and 37 to 58 years), withcells from the younger age group possessing over a two-fold greaterosteogenic potential.

[0220] Osteogenesis is defined by three phases: the proliferation ofosteogenic precursors, maturation of these precursors into osteoblasts(accompanied by matrix deposition) and a mineralization phase. Eachphase is essential and can dramatically affect the development of maturebone. The decrease in osteogenesis observed in older donors may be dueto three possibilities: 1) a decrease in PLA cell proliferation, 2) adecrease in the number of PLA-derived osteogenic precursors themselvesor 3) a decrease in osteogenic differentiation capacity. As shown inFIG. 8, PLA population doubling time increases slightly in older donorssuggesting that the proliferative capacity of older PLA cells diminisheswith age. However, this increase in population doubling time is notstatistically significant and is not likely to contribute to theage-dependent decrease in osteogenic potential.

[0221] In order to determine if a decrease in the number of osteogenicprecursors within the PLA contributed to our results, the average numberof CFU/AP⁺ colonies was determined. Colonies with AP activity areconsidered to be osteoprogenitors and have been previously used todetermine the number of osteogenic precursors and/or stem cells in bonemarrow (Owen, T A, et al., 1990 J. Cell Physiol. 143:420-430). Whileanimal studies indicate a decrease in the number of osteoprogenitors inbone marrow with advancing age (Bergman R J, et al., 1996 J. Bone Miner.Res. 11:568-577; Huibregtse, B A, et al., 2000 J. Orthop. Res.18:18-24)), conflicting results have been reported for human samples.Work by Glowacki and Rickard et al. indicate no age-related changes inbone marrow osteoprogenitor cells (Rickard, D J, et al., 1996 J. BoneMiner. Res. 11:312-324; Glowacki, J. 1995 Calcif. Tiss. Int. 56(Suppl1):S50-51)). In support of these studies, we find a small, butstatistically insignificant, change in the number of CFU/AP⁺ colonieswith age. This suggests that the observed age-dependent decrease inosteogenic potential may not be due to a drop in the number ofosteogenic precursors and/or stem cells within the PLA.

[0222] The decrease in PLA osteogenesis may be due to the loss ofosteogenic capacity. Several factors may influence the osteogeniccapacity of stem cells, including: 1.) cell-cell and cell-matrixinteractions; and 2.) growth factors and hormones. A recent study byBecerra et al. demonstrates a significant decrease in the osteogenicresponse of older Mesenchymal stem cells to demineralized bone matrix inrats (Becerra, J., et al., 1996 J. Bone Miner. Res. 11:1703-1714),suggesting age-related alterations in MSC-matrix interactions.Similarly, decreases in osteogenic potential in older donors have beencorrelated to the degradation of the extracellular matrix (Bailey, A J,et al., 1999 Calcif. Tiss. Int. 65:203-210). The microenvironmentsurrounding PLA cells may change with increasing age, altering cell-celland cell-extracellular matrix interactions that could inhibit osteogenicdifferentiation of PLA cells or favor their differentiation to anotherlineage (e.g adipogenic). Furthermore, the diminishment of osteogenicpotential in PLA cells may be due to gender. All donors in this studywere female. It is well documented that aging in the female isaccompanied by the loss of estrogen, coupled to a decrease in skeletalbone mass (Parfitt, A M 1990 in Bone, ed B K Hall, Vol. 1, 351-431, NewJersey: Caldwell; Hahn, T J 1993 in Textbook of Rheumatology, ed W NKelly, 1593-1627, New York: Saunders). Since osteocytes do notreplicate, bone remodeling and repair requires a continuous supply ofosteoblasts, the principal source of which is the bone marrow stroma.Estrogen is known to regulate the differentiation of bone marrow-derivedstem cells and decreases in circulating estrogen levels can be linked toa loss of stem cell osteogenic potential (Robinson, J A, et al., 1997Endocrinology 138:2919-2927; Ankrom, M A, et al., 1998 Biochem. J.333:787-794). Like bone marrow stem cells, the loss of osteogeniccapacity by PLA cells in older female donors may simply reflect thechanges that are associated with estrogen loss. A possibility is thatthe decrease in PLA osteogenic potential may be due to relatively smallchanges in all three factors discussed above, reflecting a generalphenomenon observed in aging women.

[0223] A reduction in osteoblast number and bone-forming activity,coupled to an increase in marrow cavity adipogenesis, contributes totype II or age-related, osteoporosis (Parfitt, A M 1990 in Bone, ed B KHall, Vol. 1, 351-431, New Jersey: Caldwell; Hahn, T J 1993 in Textbookof Rheumatology, ed W N Kelly, 1593-1627, New York: Saunders). Whilecurrent research is focusing on the role of bone marrow-derivedMesenchymal stem cells in osteoporosis, the age-related loss ofosteogenic capacity by adipose-derived PLA cells may provide researcherswith an alternate model system for the study of this disease.Furthermore, PLA cells may represent another viable cell-basedtherapeutic paradigm for the treatment of osteoporosis and othermetabolic bone disorders.

[0224] The use of stem cells for tissue engineering applications may bedramatically influenced by stem cell number, growth kinetics anddifferentiation potential. Each of these factors, in turn, may beaffected by the age of the donor. Several studies on bone marrow-derivedmesenchymal stem cells have reported alterations in MSC number,population doubling and differentiation potential with respect to donorage in both animal and human models (Lansdorp, P. M., et al., 1994 BloodCells 20:376-380; Becerra, J., et al., 1996 J. Bone Miner. Res.11:1703-1714; Bergman, R. J., et al., 1996 J. Bone Miner. Res.11:568-77; Gazit, D., et al., 1998 J. Cell Biochem. 70:478-88; Oreffo,R. O., et al., 1998 Clin. Sci (Colch.) 94:549-555; D-Ippoliot, G., etal., 1999 J. Bone Miner. Res. 14:1115-1122; Long, M. W., et al., 1999 J.Gerontol. A. Biol. Sci. Med. Soc. 54:B54-62; Huibregtse, B. A., et al.,2000 J. Orthop. Res. 18:18-24). We have characterized several PLApopulations by determining population doubling, differentiationpotential and average colony forming unit number with respect to donorage.

EXAMPLE 9

[0225] The following description provides adipose-derived stem cellswhich differentiated into chondrogenic tissue, and method for isolatingsaid stem cells.

Materials and Methods Reagents and Antibodies

[0226] Sodium acetate, bovine serum albumin (BSA), N-ethylmaleimide(NEM), 6-aminocaproic acid, phenylmethyl-sulfonyl fluoride (PMSF), andbenzamidine hydrochloride were all purchased from Sigma (St. Louis,Mo.). Monoclonal antibodies to type II collagen (clone II-4C11),chondroitin-4-sulfate, and keratan sulfate (clone 5-D-4) were purchasedfrom ICN Biomedical (Aurora, Ohio).

Lipoaspirate Processing

[0227] Human liposuction aspirates were obtained from ten healthyelective cosmetic surgery patients ranging in age from 20-55 years, andprocessed to obtain the processed lipoaspirate (PLA) cell populations.All procedures were approved by the Human Subject Protection Committee(HSPC) under protocol number HSPC #98-08-011-02. Raw lipoaspirates wereprocessed based on the method described in Example 7, supra. Briefly,the lipoaspirates were washed extensively in phosphate-buffered saline(PBS) and then incubated with 0.075% collagenase (Sigma, St. Louis, Mo.)at 37° C. for thirty minutes with gentle agitation. The collagenase wasneutralized by adding an equal volume of Dulbecco's Modified EagleMedium (DMEM, Cellgro, Herndon, Va.), and FBS, and the cellularsuspension was centrifuged at 260 g for five minutes. The resultant cellpellet was resuspended in 1% erythrocyte lysis buffer (0.16 M NH₄Cl) tolyse the contaminating reb blood cells. The cell suspension wascentrifuged at 260 g for five minutes to isolate the PLA fraction. ThePLA pellet was resuspended in control medium (DMEM, 10% FBS, and 1%antibiotics-antimycotics) and maintained at subconfluent concentrationsat 37° C. with 5% CO₂. Human foreskin fibroblasts (HFFs) were similarlyharvested through enzymatic digestion with collagenase and maintained atsubconfluent levels in control medium.

Chondrogenic Differentiation

[0228] After culture expansion to three passages (P3), the PLA cellswere trypsinized and resuspended in control medium at a concentration of10⁷ cells/ml. Chondrogenic differentiation was induced using a micromassculture protocol as previously described with some modifications(Ahrens, P B, et al., 1977 Dev. Biol. 60:69-82; Denker, A E 1995Differentiation 59:25-34). Ten microliter drops of the PLA cellularsuspension were placed in the center of each well of a 24-well tissueculture plate and on chamber slides. The cells were placed in anincubator at 37° C. at 5% CO₂ for two hours to allow cell adherence. Thepellets were gently overlaid with control medium and incubatedovernight. The medium was replaced by chondrogenic medium [DMEM with 1%FBS supplemented with 10 ng/ml TGF-β1 l (R&D Systems, Minneapolis,Minn.), 6.25 μg/ml insulin (Sigma), and 6.25 μg/ml transferrin (Sigma)].The pellets were induced for six days in chondrogenic medium. At day sixand thereafter, 50 μg/ml ascorbic acid-2-phosphate (Sigma) was added tothe chondrogenic medium mixture. PLA pellets were harvested at days two,seven, and fourteen after initial induction for analysis. In order toidentify optimal culture conditions for the induction of chondrogenicdifferentiation, PLA cells were also induced with dexamethasone (Sigma)alone at a concentration of 0.1 μM and in combination with TGF-β1. HFFcells were cultured as above under micromass and monolayer conditions asa negative control. PLA cells, incubated as monolayer cultures, did notform three-dimensional nodules and were unavailable for paraffinembedding and histologic and immunohistological analysis.

Differential Cell Density Plating

[0229] In order to assess the relationship of chondrogenic induction toPLA cell, micromass cultures were plated in chondrogenic medium at cellconcentrations of 1×10⁵, 1×10⁶, 2.5×10⁶, 5×10⁶, 1×10⁷, 2×10⁷, and 5×10⁷cells per milliliter (cells/ml). The micromass cultures were thensubjected to chondrogenic culture conditions and the onset of noduleformation noted.

Alcian Blue Staining

[0230] In order to detect the presence of highly sulfated proteoglycans,characteristic of cartilaginous matrices, induced PLA pellets werestained using Alcian blue at acidic pH (Lev, R and S Spicer 1964 J.Histochem Cytochem. 12:309). Micromass cultures were fixed with 4%paraformaldehyde in PBS for fifteen minutes, followed by a five minuteincubation in 0.1 N HCl to decrease the pH to 1. The cultures werestained overnight with 1% Alcian blue 8GX (Sigma) in 0.1 N HCl (pH 1).The cells were washed twice with 0.1 N HCl to remove nonspecificstaining and then air-dried. For paraffin sections, cellular noduleswere harvested, washed twice in PBS and fixed in 4% paraformaldehyde forone hour. The nodules were embedded in paraffin and cut intofive-micrometer sections. Paraffin sections of PLA nodules were preparedas described and stained with standard Alcian blue staining at pH 1 inorder to determine the spatial distribution of sulfated proteoglycanswithin the three-dimensional structure of the nodules. Digital imageswere acquired with a Zeiss Axioskop II microscope (Carl Zeiss, Munich,Germany) and Spot software.

Histology And Immunohistochemistry

[0231] Histologic evaluation of PLA paraffin sections was performedusing standard hematoxylin & eosin (H&E) to determine cellularmorphology and Goldner's trichrome stain to detect the presence ofcollagen in the extracellular matrix. For immunohistochemistry, paraffinsections were first deparaffinized in xylene and then hydrated indecreasing ethanol solutions (100% to 70%). To facilitate antibodyaccess to epitopes, sections were predigested for one hour at 37° C. in0.5 ml chondroitinase ABC (Sigma) in 50 mM Tris (Gibco BRL), pH 8.0, 30mM sodium acetate containing 0.5 mg/ml BSA, 10 mM NEM. The sections wereincubated in 3% H₂O₂ for fifteen minutes to quench endogenous peroxidaseactivity, followed by incubation in 10% horse serum to block nonspecificbinding. The sections were subsequently incubated for one hour at 37° C.with primary antibodies to the following: human type II collagen,chondroitin-4-sulfate, and keratan sulfate at dilutions of 1:10, 1:50,and 1:250, respectively. Incubation in normal horse serum in lieu ofmonoclonal antibodies was performed to serve as a negative control.Reactivity was detected with the Vectastain ABC kit (VectorLaboratories, Burlingame, Calif.) according to the manufacturer.

cDNA Synthesis and RT-PCR

[0232] Total RNA was isolated from untreated PLA cells, PLA nodules, andHFFs. Briefly, RNA was isolated using the following method (RNA-Easy,Qiagen). The RNA was used for oligo dT-primed cDNA synthesis usingMMLV-RT enzyme (Promega). Equivalent amounts of cDNA were subjected toPCR amplification using primer pairs designed to: human type I collagenα1 chain (CN I), human type II collagen α1 chain (CN II), human type Xcollagen α1 chain (CN X), human large aggregating proteoglycan oraggrecan (AG) and human osteocalcin (OC). The primer pairs used wereobtained from published GeneBank sequences (Table 4) and are as follows:TABLE 4: Expected Product Gene accession # Primer #1 Primer #2 Size CN INM_(—000088) 5′-CAT CTC CCC 5′-CTG TGG AGG AGG 598 bp TTC GTT TTT GA-GTT TCA GA-3′ 3′ CN II Published 5′-CTG CTC GTC 5′-AAG GGT CCC AGG IIA*:432 bp (148) GCC GCT GTC TTC TCC ATC-3′ IIB*: 225 bp CTT-3′ N XNM_000493 5′-TGG AGT GGG 5′-GTC CTC CAA CTC 601 bp AAA AAG AGG CAG GATCA-3′ TG-3′ AG X17406 5′-GCA GAG ACG 5′-GGT AAT TGC AGG 504 bp CAT CTAGAA GAA CAT CAT T-3′ ATT G-3′ OC X04143 5′-GCT CTA GAA 5′-GCG ATA TCCTAG 310 bp TGG CCC TCA ACC GGG CCG TAG-3′ CAC TC-3′

[0233] Primer pairs for type II collagen, type X collagen and aggrecanwere confirmed against articular cartilage samples as a positivecontrol. Calculated optimal annealing temperatures (OLIGO PrimerAnalysis Software, National Biosciences Inc., Plymouth, Minn.) were usedfor each primer pair. Templates were amplified for 35 cycles and the PCRproducts were analyzed using conventional agarose gel electrophoresis.

Effect Of Passage On The Chondrogenic Potential Of PLA Cells

[0234] To examine the effect of multiple cell passaging on thechondrogenic potential of human PLA cells, monolayer cultures werepassaged fifteen times, with cell fractions taken at the first, thirdand fifteenth passages. The cell fractions were placed in micromasscultures, grown in chondrogenic medium and chondrogenic differentiationwas assessed by Alcian blue staining.

PLA Clonal Isolation

[0235] Freshly isolated PLA cells were plated out at a density of 100cells per 100 mm² tissue culture dish, to promote the formation ofcolonies from single cells. Cultures were expanded in control mediumuntil the appearance of distinct colonies. Colonies derived from singlePLA cells were isolated using sterile cloning rings, then harvested with0.25% trypsin digestion. The dissociated cells were seeded into 24-wellplates and expanded. PLA clones were induced toward the chondrogeniclineage as described above and chondrogenic differentiation wasconfirmed by Alcian blue staining and type II collagenimmunohistochemistry.

Results

[0236] Human lipoaspirates were processed to obtain the PLA cellpopulation. The PLA was placed into high-density micromass culturessupplemented with TGF-β1, insulin, transferrin, and ascorbic acid toinduce chondrogenic differentiation. Chondrogenesis was assessedhistologically at two, seven, and fourteen days using standardhistologic assays. In addition, immunohistochemistry was performed withantibodies to type II collagen, chondroitin-4-sulfate, and keratansulfate. Finally, RT-PCR analysis was performed to confirm theexpression of type I, type II, and type X collagen as well ascartilage-specific proteoglycan and aggrecan.

[0237] All TGF-β1-treated micromass cultures formed three-dimensionalspheroids within 48 hours of induction that stained positively withAlcian blue, suggestive of cartilaginous nodule formation.Immunohistochemistry confirmed the presence of type II collagen,chondroitin-4-sulfate, and keratan sulfate throughout the extracellularmatrix of the nodules. Finally, RT-PCR analysis confirmed the expressionof cartilage-specific type II collagen, aggrecan, and cartilage-specificproteoglycan.

PLA Cells Form Chondrogenic Nodules

[0238] Pre-cartilage mesenchymal cells and multi-lineage stem cells canbe induced toward the chondrogenic lineage using a high-densitymicromass culture technique, followed by induction with pro-chondrogenicfactors (Ahrens, P B, et al., 1977 Dev. Biol. 60:69-82; Denker, A E, etal., 1995 Differentiation 59:25-34; Johnstone, B, et al., 1998 Exp. CellRes. 238:265-272). Consistent with these studies, human ProcessedLipoaspirate (PLA) cells, cultured under high-density micromassconditions and induced with chondrogenic medium, containing transforminggrowth factor-beta 1 (TGF-β1), insulin, and transferrin, condensed intothree-dimensional spheroids as early as twenty-four hourspost-induction. At this time period, the PLA nodules were visible to thenaked eye as white, round structures measuring approximately 1-2 mm indiameter. Nodules formed in 100% of over 500 treated micromass cultures.Small spheroids formed in untreated micromass cultures occasionally(10%) and may be an effect of the culture conditions themselves. No PLAnodules were observed in TGF-β1 -treated or untreated PLA monolayercultures. PLA nodules became larger in size with culture time andsmaller adjacent nodules could be visualized under a microscope afterseven days in culture. In some cases, adjacent PLA nodules coalescedinto a larger, cellular aggregates with increased culture time and isconsistent with the proposed cellular interactions and recruitment thatare essential to chondrogenesis (Ahrens, P B, et al., 1977 Dev. Biol.60:69-82).

[0239] In order to assess the effect of cell number on PLA noduleformation, differential plating studies were performed. No evidence ofspheroid formation was seen in cultures plated at a density of less than5×10⁶ cells/ml. PLA cells plated at increasing densities (i.e. above1×10⁷ cells/ml) underwent nodule formation more rapidly and, in somecases, were more likely to undergo spheroid formation in the absence ofTGF-β1. The addition of dexamethasone to chondrogenic medium, containingTGF-β1, has been shown to lead to the formation of larger cartilaginousaggregates (Johnstone, B., et al., 1998 Exp. Cell Res. 238:265-272).Consistent with this, the addition of dexamethasone resulted in largerspheroids when compared to nodules formed with TGF-β1 stimulation.Cultures treated with dexamethasone alone did not form nodules,suggesting that TGF-β1 is crucial to nodule formation by PLA cells.Finally, no evidence of nodule formation was observed in micromass andmonolayer HFF cultures treated with chondrogenic medium, confirming thespecificity of our chondrogenic conditions.

PLA Nodules Contain an Extracellular Matrix Rich in SulfatedProteoglycans

[0240] Cartilagenous matrices contain very high quantities ofpolyanionic sulfated glycoasminoglycans (GAGs), such as chondroitin 4-and 6-sulfate, and are characterized by the ability to stain positivelywith Alcian blue at low pH (R Lev and S Spicer 1964 J. Histochem.Cytochem. 12:309). In order to confirm the cartilaginous nature of thePLA nodules, histologic analysis was performed on whole-mount PLAnodules, plated on chamber slides, and paraffin sections. Initialtreatment of PLA cultures with chondrogenic medium resulted in cellularcondensation within 24 hours (FIG. 12, Panel A). Condensing PLA cellsexhibited a low level of Alcian Blue staining, suggesting the initialformation of a sulfated extracellular matrix. PLA condensation wasfollowed by ridge formation and increased staining by Alcian Blue,indicating an increase in matrix secretion (Panel B). Intense AlcianBlue staining and spheroid formation was observed after 48 hourspost-induction (Panel C). In contrast, untreated PLA cells in micromasscultures did not show any regions of positive Alcian blue staining(Panel D).

[0241] In addition to whole-mount PLA samples, paraffin sections of PLAnodules were prepared in order to assess the three-dimensionalarchitecture of the nodule. The morphology of the paraffin-embeddedsections, as analyzed by hematoxylin and eosin staining, showed a flat,peripheral layer of fibroblast-like cells that resembled perichondralcells, surrounding an inner core of rounder cells at two dayspost-induction (FIG. 13, Panel A). After fourteen days of treatment,nodules became more hypocellular with increasing deposits ofextracellular matrix into the core (Panel B). Goldner's trichromestaining, which indicates the presence of collagenous matrix (greencolor), confirmed the H&E pattern (Panels C and D). Faint backgroundlevels of collagenous matrix were observed in the nodule sections at twodays (Panel C), compared with higher levels of collagen seen in thenodule core at fourteen days post-induction (Panel D). Alcian bluestaining of the paraffin sections was similar to the whole-mountpreparations, confirming the formation of cartilaginous matrix rich insulfated proteoglycans after two days induction (Panel E). Increasedstaining intensity in the central core region was observed at fourteendays post-induction (Panel F), suggesting an increased secretion ofsulfated proteoglycans as the cells mature down the chondrocyticpathway. In summary, our histological staining results confirm theformation of cartilage-like PLA nodules, associated with anextracellular matrix rich in collagens and sulfated proteoglycans.

PLA Nodules Express Cartilage-Specific Proteins

[0242] Immunohistochemical analysis was used to detect the presence oftype II collagen, an extracellular matrix component highly specific forcartilaginous tissue, and chondroitin-4-sulfate and keratan sulfate, twoof the main monomeric components of cartilage proteoglycans. After twodays induction, areas of strong immunoreactivity tochondroitin-4-sulfate and keratan sulfate were seen along the outerperiphery of the spheroids and throughout the core and is supportive ofour Alcian Blue staining results (FIG. 14, Panels A and C). Asignificant increase in chondroitin-4-sulfate and keratan sulfateexpression within the nodule core was noted over the course of two weeks(Panels B and D). In contrast, positive type II collagenimmunoreactivity was not evident in the PLA nodules at day two (PanelE). Rather, collagen type II expression appeared at day sevenpost-induction, with strong expression appearing at day fourteen (PanelF). Whole-mount cultures of TGF-β1-treated PLA micromass cultures alsoshowed intense type II collagen reactivity while untreated micromass PLAcultures showed no staining . In addition, no staining was observed inparaffin sections incubated in normal horse serum instead of primarymonoclonal antibodies, supporting the specificity of the type IIcollagen, chondroitin-4-sulfate, and keratan sulfate antibodies. Takentogether, the immunohistochemical results support the histologicalstaining data and suggest the presence of a cartilaginous matrix in PLAnodules.

Chondrogenic Differentiation of Single-Cell Derived Clonal Populations

[0243] The apparent chondrogenic differentiation by PLA cells may resultfrom contamination of the lipoaspirate by pre-chondrogenic cells ratherfrom the presence of a multipotential cell. Therefore to determine ifour results are due to differentiation of multipotential PLA cells, weisolated and confirmed the multilineage potential of single-cell derivedPLA clones. PLA clonal populations (i.e. adipo-derived mesodermal stemcells or ADSCs) demonstrated the ability to undergo chondrogenicdifferentiation in addition to osteogenic and adipogenicdifferentiation. PLA clonal populations induced toward the osteogenicand adipogenic lineages exhibited classic lineage-specific histologicalmarkers (alkaline phosphatase activity-osteogenesis; Oil-Red-Oaccumulation-adipogenesis) (unpublished data). Like the heterogeneousPLA cultures, PLA clonal populations also underwent spheroid formationwithin forty-eight hours of induction in chondrogenic medium. Inaddition, the PLA nodules secreted an extracellular matrix rich in typeII collagen and highly sulfated proteoglycans.

PLA Cells Retain Chondrogenic Potential After Extended Culture

[0244] Culture time and passage number can affect the differentiativecapacity of many cell types. To assess the effect of passaging on thechondrogenic potential of PLA cells, PLA cells were passaged inmonolayer cultures as many as fifteen times (175 culture days) andcultured under high-density conditions to induced chondrogenesis. PLAcells retained their chondrogenic differentiation potential throughoutthis extended culture period, as evidenced by their ability to formthree-dimensional spheroids after induction with chondrogenic medium.Finally, both early and late passage PLA nodules secreted anextracellular matrix rich in highly-sulfated proteoglycans as evidencedby the positive staining with Alcian blue (FIG. 22). Cellular nodulesfrom all culture passages (i.e. P1 to P15) had a very similarappearance: a flat, peripheral layer of fibroblast-like cells resemblingthe perichondrium surrounding an inner core of rounder cells.

RT-PCR Analysis Confirms the Expression Of Cartilage-Specific Collagens

[0245] RT-PCR analysis of PLA nodules was performed using primersspecific to the genes for human type I collagen, type II collagen, andtype X collagen, as well as aggrecan and osteocalcin. Untreated HFF andhuman PLA cells cultured under micromass conditions were analyzed asnegative controls. RT-PCR analysis of PLA nodules confirmed theexpression of type II collagenα1 (CN II) at day 7 and day 14 only (FIG.15). Moreover, decrease in CN II expression was observed between 7 and14 days induction. Both splice variants of CN II (IIA and IIB—type IIBvariant shown) were observed at both time points.

[0246] In contrast to day seven and fourteen nodules, CN II expressionwas not observed in 2-day nodules, confirming our immunohistochemicaldata. As expected, CN II was not observed in HFF micromass cultures .However, small amounts of CN II mRNA were present in the untreated PLAcells. Chondrogenic differentiation was further confirmed by examiningnodules for the expression of the large aggregating proteoglycan, oraggrecan. Aggrecan has been shown to be specific to cartilage andaccumulates at the onset of over chondrogenesis (Kosher, R A, et al.,1986 J. Cell Biol. 102:1151-1156). Aggrecan expression was observed atboth 2 and 7 days induction and was absent in 14 day PLA nodules.Aggrecan expression was specific to treated PLA nodules, as noexpression was noted in control PLA cells or in HFF cultures.

[0247] Further characterization of PLA nodules was performed byassessing the expression of the α1 chains of type I and type X collagen.Collagen type I expression is known to be up-regulated in osseoustissues and is down-regulated during chondrogenic differentiation(Kosher, R A, et al., 1986 J. Cell Biol. 102:1151-1156; Shukunami, C.,et al., 1998 Exp. Cell Res. 241:1-11). Consistent with this, CN Iexpression was observed in 2-day treated PLA nodules only. Similar to CNII, low levels of CN I were observed in untreated PLA cells, suggestingthat undifferentiated PLA cells are associated with a collagenous matrixthat is dramatically remodeled as differentiation proceeds. CN Xexpression was not observed in PLA nodules at two and seven dayspost-induction but appeared at the 14-day time point. No CN X wasobserved in untreated PLA cells or in HFF controls. Collagen type X isspecific to hypertrophic chondrocytes and may signal the progression toendochondral ossification and bone formation (Linsenmayer, T F, et al.,1988 Pathol. Immunopathol. Res. 7:14).

[0248] To confirm the absence of ossification and bone formation withinthe PLA nodules, RT-PCR analysis was performed using primers toosteocalcin, a bone-specific gene (Price P A 1989 Connect. Tissue Res.21:51-57). As expected, osteocalcin expression was absent in all treatedand untreated PLA samples. Taken together, the expression ofcartilage-specific aggrecan, both type II and X collagen, together withthe decreased expression of type I collagen supports the chondrogenicdifferentiation by PLA cells.

Discussion

[0249] The repair of cartilaginous defects remains a significantclinical challenge. Damaged articular cartilage has a limited potentialfor repair and large defects do not heal spontaneously. When the damageextends into the subchondral bone, the repair process is sporadic andthe original articular cartilage is replaced by fibrocartilage and scartissue, which are structurally inferior to the hyaline architecture ofnormal articular cartilage.

[0250] Conventional treatment modalities for cartilage defects includemarrow stimulation techniques (e.g. subchondral drilling) and jointarthroplasty (I H Beiser and O I Knat 1990 J. Foot Surg. 29:595-601;Gilbert, J E 1998 Am. J. Knee Surg. 11:42-46; T Minas and S Nehrer 1997Orthopedics 20:525-538; O'Driscoll, S W 1998 J. Bone Joint Surg. Am.80:1795-1812). More recently, newer strategies have been developed, suchas the use of osteochondral, perichondral, and periosteal allografts(Bouwmeester, S J, et al., 1997 Int. Orthop. 21:313-317;Carranza-Bencano, A, et al., 1999 Calcif. Tissue Int. 65:402-407;Ghazavi, M T, et al., 1997 J. Bone Joint Surg. Br. 79:1008-1013;Homminga, G N, et al., 1990 J. Bone Joint Surg. Br. 72:1003-1007).Unfortunately, these options do not result in complete regeneration ofthe original hyaline architecture. More importantly, the joint is notcapable of normal weight-bearing and physical activity over prolongedperiods of time.

[0251] Cell-based tissue engineering strategies represent a promisingalternative to conventional techniques. First-generation tissueengineering strategies are currently employed clinically usingautologous chondrocyte implantation (Brittberg, M., et al., 1994 NewEngl. J. Med. 331:889-95; Chen, F S, et al., 1997 Am. J. Orthop.26:396-406; Gilbert J E 1998 Am. J. Knee Surg. 11:42-46; Richardson J B,et al., 1999 J. Bone Joint Surg. Br. 81:1064-1068). However, limitedavailability of donor sites for chondrocyte harvest, the requirement forlengthy in vitro culture expansion, and donor site morbidity limit thepracticality of this technique. It is important to identify othersources of chondrocytic precursors.

[0252] Several cell types have been shown to undergo in vitro and invivo chondrogenesis, including rat calvarial clonal cell lines andprimary cells, the murine embryonic C3H10T1/2 cells, andperiosteum-derived and bone marrow-derived precursors from severalanimals including rabbits, rats, horses, and goats (Denker, A E, et al.,1995 Differentiation 59:25-34; Fortier, L A, et al., 1998 Am J. Vet.Res. 59:1182-1187; Grigoriadis, et al., 1996 Differentiation 60:299-307;Grigoriadis, et al., 1988 J. Cell. Biol. 30 106:2139-2151; Iwasaki, etal., 1995 J. Bone Joint Surg. Am. 77:543-554; Johnstone, et al., 1998Exp. Cell Res. 238:265-272; Nakahara, et al., 1990 Bone 11:181-188;Shukunami, et al., 1996 J. Cell. Biol. 133:457-468). However, thereremains a large potential reservoir of osteochondrogenic precursors fromother tissue types that have yet to be studied. The interconversionability of various mesodermal cell types has been reported in manystudies. Specifically, both mature human adipocytes and adipocytesisolated from bone marrow exhibit the potential to differentiate intobone (Bennett, J H, et al., 1991 J. Cell Sci. 99(Ptl):131-139; Park, etal., 1999 Bone 24:549-554). In addition, osteoblasts transdifferentiateinto chondrocytes and muscle cells are capable of commitment to thecartilage lineage (Manduca, et al., 1992 Eur. J. Cell Biol. 57:193-201;Nathanson, M A 1985 Clin. Orthop. 200:142-158; Sampath, et al., 1984Proc. Natl. Acad. Sci. USA 81:3419-3423).

[0253] The presence of mesenchymal stem cells capable ofosteochondrogenic differentiation in human bone marrow has beenwell-documented (Mackay, et al., 1998 Tissue Eng. 4:414-428; Pittinger,et al., 1999 Science 284:143-147; Yoo, et al., 1998 J. Bone Joint Surg.Am. 80:1745-1757). Some of the advantages of using mesenchymal stemcells include their ability to proliferate rapidly in culture, theirability to differentiate into chondrogenic cells even after multiplepassages, their regenerative capacity, and a broad range of resultantchondrogenic cell types (i.e. prechondroctyes, mature chondrocytes, andhypertrophic chondrocytes). Researchers have anticipated that thedifferentiated chondrogenic tissue derived from stem cells will moreclosely resemble that seen in developing embryonic limb buds. Moreover,chondrocytes proliferate poorly in culture, are difficult to maintain,and dedifferentiate when expanded in monolayer cultures (von der Mark,et al., 1977 Nature 267:531-532). The use of autologous stem cells inplace of harvested chondrocytes in tissue engineering may be a moreefficacious alternative in the future for treatment of cartilagedefects. Unfortunately, the limited availability of donor sites and thediscomfort and pain associated with bone marrow procurement remain aconcern.

[0254] The presence of a multipotential cell population within adiposetissue, capable of differentiation into several mesenchymal tissues maybe an important finding. Adipose tissue is available in large quantitiesand relatively easy to obtain. Moreover, liposuction procedures haveminimal donor site morbidity and patient discomfort. Because of thesepractical advantages as a cell source, we sought to determine if PLAcells, like bone marrow- and periosteum-derived mesenchymal stem cells,represent a cell population with the ability to undergo chondrogenicdifferentiation.

[0255] We have confirmed the chondrogenic potential of multilineagehuman processed lipoaspirate (PLA) cells. Human PLA cells inhigh-density micromass cultures treated with TGF-β1 resulted in theformation of three-dimensional cellular nodules with cartilaginouscharacteristics. The chondrogenic nature of the differentiated cells wassupported by several findings: 1) whole-mount PLA nodules and histologicsections stained positively with Alcian blue, 2) H&E morphologyrevealing a perichondral border of cells surrounding a hypocellularchondrogenic core, 3) a collagen-rich extracellular matrix as shown byGoldner's trichrome staining, 4) expression of type II collagen,chondroitin-4-sulfate, and keratan sulfate as confirmed byimmunohistochemistry, and 5) expression of collagen type II as well ascartilage-specific aggrecan as shown by RT-PCR.

[0256] One of the earliest features of cartilage development in vivo isthe formation of cellular condensations that represent skeletalprimordia. Cartilage initially differentiates in the center of thesecondensations and is followed by a period in which the cells secrete andare surrounded by a characteristic extracellular matrix. Similar to thissituation, chondrogenic differentiation in vitro is characterized by theformation of multi-layered cellular aggregates, called spheroids ornodules. Primary nodule formation is followed by ridge formation, theaccumulation of matrix and the recruitment of adjacent cells, resultingin the expansion of the original nodule (Ahrens, et al., 1977 Dev. Biol.60:69-82; Denker, et al., 1995 Differentiation 59:25-34; Stott, et al.,1999 J. Cell Physiol. 180:314-324; Tacchetti, et al., 1992 Exp. CellRes. 200:26-33; Tavella, et al., 1994 Exp. Cell Res. 215:354-362).Consistent with these studies, PLA cells began condensing withintwenty-four hours induction with TGF-β1-containing chondrogenic mediumand formed well-defined three-dimensional spheroids by forty-eight hourspost-induction. The appearance of smaller adjacent nodules in additionto the original cartilage nodule was noted after cultures were treatedfor extended periods in chondrogenic medium, suggesting the presence offurther chondrogenic induction through possible paracrine growth factorsignaling by the maturing cartilaginous nodule. PLA nodule formation wasevident only in micromass cultures plated at a cell density higher than5×10⁶ cells/ml, consistent with previous studies describing the highcell density requirement for chondrogenesis (Rodgers, et al., 1989 CellDiffer. Dev. 28:179-187; Tsonis and Goetinck 1990 Exp. Cell Res.190:247-253 ).

[0257] Cartilage is comprised of a mixture of collagen fibrils andproteoglycans that give the tissue high tensile strength and internalswelling pressure. The predominant collagen of cartilage is collagentype II. Although this collagen is not specific to cartilage it ishighly characteristic of this tissue, as collagen type II is produced bya limited number of non-chondrogenic cell types. Positive staining usinga Goldner Trichrome stain, specific for collagens in general, confirmedthese proteins within the PLA nodule after both 2 and 14 days inductionwith chondrogenic medium. Specifically, PLA nodules treated with TGF-β1for 48 hours were associated with an extracellular matrix containing lowlevels of collagen type II, suggesting that PLA cells have undergonepreliminary chondrogenic differentiation. Collagen type II levelsappeared to increase with induction time. In addition to collagen typeII, cartilagenous matrices also contain high levels of sulfated GAGs,such as chondroitin-4- and -6-sulfate that are typically associated withproteoglycans such as aggrecan. Consistent with this, histologicalstaining with Alcian Blue confirmed the presence of sulfatedproteoglycans as early as 24 hours induction, increasing as the PLAnodule became more defined (i.e. 2 days). Increased Alcian Blue stainingwas also observed as far as 14 days induction, localizing to the nodulecore and surrounding individual cells. Similar results were alsoobserved when nodules were stained with antibodies specific to keratan-and chondroitin-sulfate confirmed and immunohistochemical studiesconfirmed the presence of these components and further supports thepresence of chondrogenic cells within the PLA nodule.

[0258] In support of our immunohistochemical results, RT-PCR analysisconfirmed the expression of CN II in PLA nodules induced for 7 and 14days, with a lower level of this gene being observed at day 14. No CN IIexpression was observed after 48 hours induction with chondrogenicmedium. The expression of CN II in PLA nodules is supportive of thechondrogenic phenotype. Our immunohistochemical results showed asignificant level of both chondroitin- and keratan-sulfate specificallyin induced PLA nodules. It is known that chondroitin-4- and 6- sulfateare the main monomeric components of the cartilage-specific protein,aggrecan (Hall, B K 1981 Histochem. J. 13:599-614). Aggrecan has beenshown to be cartilage-specific and accumulates at the onset of overtchondrogenesis, coincident with cellular condensation (Kosher, et al.,1986 J. Cell Biol. 102:1151-1156).

[0259] We confirmed the chondrogenic nature of the PLA nodule byassessing the expression of aggrecan. As shown in FIG. 22, theexpression pattern of aggrecan overlapped with that of CN II at day 7.In addition, the expression of aggrecan preceded that of CN II. However,in contrast to CN II, aggrecan was not observed in PLA nodules inducedfor 14 days. Aggrecan is a cartilage-specific protein that consists of amultidomain protein core containing binding sites for sulfatedproteoglycans (Hardinghamn, et la., 1984 Prog. Clin. Biol. Res.151:17-29). The primers used to detect aggrecan in this study weredesigned to the C-terminus, which contains the G3 globular domain, asite that undergoes alternative splicing and is proteolytically cleavedin mature cartilage (Fulop, et al., 1993 J. Biol Chem. 268:17377-17383).The absence of aggrecan in day 14 PLA nodules may therefore represent analternatively spliced form of aggrecan that lacks the C-terminus.However, no aggrecan at day 14 was observed when RT-PCR was performedusing primers designed to the N-terminus, suggesting that aggrecan is nolonger expressed after two weeks induction with chondrogenic medium.

[0260] In addition to aggrecan and CN II, PLA nodules expressed bothtype I and type X collagen at distinct time points. Day 2 PLA noduleswere characterized by the expression of both CN I and CN II. However, incontrast to aggrecan and CN II, the expression pattern of CN I washighly restricted and did not appear beyond the two day time point.Interestingly, low levels of both CN I and CN II were observed inuntreated PLA control cells. Consistent with this, both type I and typeII collagen mRNA have been found in many developing embryonic tissuesand basal levels of these collagen subtypes can be detected inpre-cartilage mesenchymal precursors prior to chondrogenicdifferentiation (Lisenmeyer, et al., 1973 Dev. Biol. 35:232-239; Dessau,et al., 1980 J. Embryol. Exp. Morphol. 57:51-60; Cheah, et al., 1991Development 111:945-953; Kosher and Solursh 1989 Dev. Biol 131:558-566;Poliard, et al., 1995 J. Cell Biol. 130:1461-1472). Finally, thedecrease in CN II expression in day fourteen nodules coincided with theappearance of CN X, a collagen indicative of hypertrophic chondrocytes(Kirsch, et al., 1992 Bone Miner. 18:107-117; Linsenmayer, et al., 1988Pathol. Immunopathol. Res. 7:14-19).

[0261] The appearance of collagen type X and the hypertrophic phenotypemay precede possible nodule ossification and bone formation. However,PLA nodules did not express osteocalcin, a bone-specific gene expressedin cells differentiating toward the osteogenic lineage (Price, et al.,1983 Biochem. Biophys. Res. Commun. 117:765-771). Despite the expressionof collagen type X, mature hypertrophic chondrocytes with theircharacteristic lacunae were not seen in PLA nodules. However, a similarresult has been described by Denker et al. when C3H10T1/2 murinepluripotent cells were placed in micromass cultures and treated withTGF-β1 (Denker, et al., 1995 Differentiation 59:25-34). Hypertrophicchondrocytes were only observed in place of nodules when cultures weretreated with BMP-2 (139). It therefore may be necessary to induce PLAmicromass cultures with BMP-2 to fully induce hypertrophy and induce theformation of mature chondrocytes.

[0262] Taken together, our histologic, immunohistochemical, and RT-PCRdata support the differentiation of PLA cells toward the chondrogeniclineage. However, the processed lipoaspirate is a heterogeneouspopulation of cells and may contain several cell types of variousmesodermal lineages. Specifically, there may exist chondrogenicprecursors in the lipoaspirate that are capable of spontaneousdifferentiation, as well as a subpopulation of multipotential cells(i.e. PLA stem cells). The isolation of PLA clones derived from singlePLA cells and their multilineage differentiation (chondrogenesis,osteogeneis, and adipogenesis) supports the presence of multipotentialstem cells (adipo-derived mesodermal stem cells) within thisheterogeneous cell population.

[0263] In order to apply cell-based tissue engineering techniques to theclinical setting, a number of criteria must be met. The cell populationused as the cellular vehicle should be abundant and easy to obtain,expandable in tissue culture, able to maintain its differentiativeability through multiple passages, and exhibit properties equivalent tothe native target tissue. The healing of articular cartilage defectsusing stem cells harvested from bone marrow has been successfullyreported in various animal models (Angele, et al., 1999 Tissue Eng.5:545-554; Butnariu-Ephrat, M., et al.,1996 Clin. Orthop. 330:234-43;Wakitani, et al., 1994 J. Bone Joint Surg. Am. 76:579-592). However, thebone marrow harvest is painful and yields low number of stem cells forclinical use, usually requiring in vitro expansion. Adipose tissue isplentiful and easy to obtain with relatively minimal discomfort. PLAcells can be harvested from a relatively small amount of adipose tissuein large numbers (Zuk, P., et al., 2001 Tissue Engineering 7:209-226),thereby, obviating the need for lengthy culture expansions. Whileelective cosmetic surgery is the most common source of lipoaspirates,sufficient adipose tissue could also be obtained through a small-borecannula for non-cosmetic surgery patients requiring reconstruction,making this technique available to a wide variety of patients. In thisexample, the chondrogenic capacity of multipotential adipose-derivedstem cells is demonstrated and shows that the stem cells retain theirability to differentiate even after long-term culture. Finally,adipose-derived stem cell nodules exhibit many properties consistentwith native cartilage tissue.

[0264] The fulfillment of these properties, together with the potentialabundance of PLA cells, make these multipotential cells an ideal systemfor tissue engineering strategies. In addition, these cells may beappropriate for the study of chondrogenesis in both in vitro culturestudies and in vivo animal models. The identification of chondrogenicprecursors has important implications for the repair of articularcartilage defects. The abundance and easy accessibility of adiposetissue makes it a feasible alternative for cartilage reconstruction(Asahina, et al., 1996 Exp. Cell Res. 222:38-47; Atkinson, et al., 1997J. Cell Biochem. 65:325-339; Chimal-Monroy and Diaz de Leon 1999 Int. J.Dev. Biol. 43:59-67; Denker, et al., 1999 Differentiation 64:67-76;Klein-Nulend, et al., 1998 Tissue Eng. 4:305-313; Martin, et al., 1999Exp. Cell Res. 253:681-688; Quarto, et al., 1997 Endocrinology138:4966-4976; Shukunami, et al., 1998 Exp. Cell Res. 241:1-11).

EXAMPLE 10

[0265] The following description provides methods for isolating stemcells from adipose tissues, where the stem cells differentiate intomyogenic tissue.

Methods Differentiation and Tissue Culture Reagents

[0266] Hydrocortisone, collagenase and paraformaldehyde were purchasedfrom Sigma (St. Louis, Mo.). Horse Serum (HS) was purchased from LifeTechnologies (Grand Island, N.Y.). Phospho-Buffered Saline (PBS), 0.25%trypsin/l mM EDTA (trypsin/EDTA), Dulbecco's Modified Eagle's Medium(DMEM) and antibiotic/antimycotic solution were purchased from CellGro(Herndon, Va.). Fetal Bovine Serum (FBS) was purchased from Hyclone(Logan, Utah).

PLA Preparation and Culture

[0267] Human adipose tissue, obtained from eight patients (mean age=39.3years, range 25-58 years) undergoing elective Suction-Assisted Lipectomy(SAL), according to patient consent protocol HSPC #98-08 011-02(University of California Los Angeles) was processed as described,according to the method described in Example 7, supra, to obtain theProcessed Lipoaspirate (PLA) cell population. Briefly, the rawliposuctioned aspirates were washed extensively with sterile PBS inorder to remove blood cells, saline and local anesthetics. Theextracellular matrix was digested with 0.075% collagenase 37° C. for 30minutes to release the cellular fraction. Collagenase was inactivatedwith an equal volume of DMEM containing 10% FBS. The infranatant wascentrifuged at 250×g for 10 minutes to obtain a high-density PLA cellpellet. The pellet was resuspended in DMEM/10% FBS and an ErythrocyteLysis Buffer (0.16M NH₄Cl) was added for 10 minutes to lysecontaminating erythrocytes. Following an additional centrifugation step,the PLA cell pellet was resuspended in DMEM/10% FBS and plated in 100 mmtissue culture dishes at a density of 1×10⁶ cells per plate. PLA cellswere maintained in Control Medium (CM—DMEM, 10% FBS, 1%antibiotic/antimycotic) at 37° C. and 5% CO₂. The culture medium waschanged twice weekly. Confluent PLA cultures (approximately 80%confluence) were passaged at a ratio of 1:3 in trypsin/EDTA. For controlstudies, a human foreskin fibroblast cell line, HFF (American TypeCulture Collection, Manassas, Va.) and a human skeletal muscle cellline, SKM (Clonetics, Walkersville, Md.) were maintained at 37° C./5%CO₂ in CM and a myogenic maintenance medium (SKM—Clonetics),respectively.

Myogenic Differentiation

[0268] To induce optimal myogenesis, PLA cells were plated at a densityof 1×10⁴ cells onto 35 mm tissue culture dishes and incubated overnightin CM to allow adherence. Optimal myogenesis was obtained by incubatingPLA cells in Myogenic Medium (MM=CM supplemented with 5% Horse Serum and50 μm hydrocortisone to promote proliferation, a key event in myogenicdifferentiation) (196). PLA cells were induced in MM for 1, 3 and 6weeks. Medium was changed twice weekly until the experiment wasterminated. SKM and HFF cells were induced for 1, 3 and 6 weeks in MM aspositive and negative controls, respectively.

Immunohistochemistry

[0269] To assess myogenic differentiation, PLA cells were seeded onto8-well chamber slides at a density of 5×10³ cells per well and allowedto adhere in CM overnight. Cells were induced in MM for 1, 3 and 6weeks. Following induction, the cells were rinsed twice with PBS andfixed with 4% paraformaldehyde for 20 minutes at 4° C. The cells wereincubated with 3% hydrogen peroxide for 5 minutes to quench endogenousperoxidase activity. Non-specific epitopes were blocked by a 30 minuteincubation in Blocking Buffer (BB; PBS, 1% HS, 0.1% Triton X-100). Thecells were incubated at 4° C. overnight with either a monoclonalantibody to human MyoD 1 (Dako; Carpenteria, Calif.) or monoclonalantibody to human fast twitch skeletal muscle myosin heavy chain(Biomeda Corp.; Foster City, Calif.). Following incubation, the cellswere washed with BB and incubated at room temperature for 2 hours in BBcontaining a horse anti-mouse IgG secondary antibody conjugated tobiotin. The secondary antibody was visualized using the VectaStain ABCkit (Vector Labs; Burlingame, Calif.) according to manufacturer'sspecifications. The cells were counterstained with hematoxylin for 3minutes. SKM cells induced in MM were processed as above as a positivecontrol. PLA cells cultured in CM and HFF cells induced in MM wereanalyzed as negative controls.

RT-PCR Analysis

[0270] Total RNA was isolated from PLA cells treated with MM for 1, 3and 6 weeks. RNA was isolated according to the method described inExample 9 above. Five micrograms (5 ug) of total RNA was used for oligodT-primed cDNA synthesis using Murine Maloney Leukemia Virus ReverseTranscriptase (MMLV-RT; Promega; Madison, Wis.). The resulting cDNA wasused as a template for PCR analysis using primer pairs designed to humanMyoD1 (Accession; NM_(—)002478) and human skeletal muscle myosin heavychain (Accession; X03740). The primer pairs used and the expected PCRproduct sizes were as follows: MyoD1: 5′-AAGCGCCATCTCTTGAGGTA-3′(forward primer) and 5′-GCGCCTTTATTTTGATCACC-3′ (reverse primer); 500bp; myosin heavy chain: 5′-TGTGAATGCCAAATGTGCTT-3′ (forward primer) and5′-GTGGAGCTGGGTATCCTTGA-3′ (reverse primer); 750 bp. MyoD1 and myosinwere amplified using Taq polymerase (Promega) for 35 cycles in a totalreaction volume of 100 ul. Duplicate reactions were performed usingprimers designed to the housekeeping gene, β-actin, as an internalcontrol. PCR products were resolved by agarose gel electrophoresis. PCRamplification of cDNA obtained from PLA cells cultured in CM and HFFcells induced in MM was performed as negative controls.

Immunohistochemical Quantification And Data Analysis

[0271] To quantitate myogenesis, a total of five hundred PLA cells fromeach induction time point were manually counted at 200×magnificationusing an “Axioskop 2” inverted microscope (Carl Zeiss Inc; Thornwood,N.Y.) and the number of MyoD1 and myosin positive cells determined. Thenumber of MyoD1 and myosin-positive cells was expressed as a percentageof the total 500 cells (% total PLA cells) and was used as an indicationof the degree of myogenic differentiation. All studies were performed oneight patients and the mean number of MyoD1 and myosin-positive cellscalculated, together with the standard error of the mean (±SEM).Myogenic differentiation in both the experimental and control groupsdescribed above was analyzed for statistical significance using aone-way analysis of variance (ANOVA). A p value of less than 0.05 wasconsidered significant.

Results Induced Stem Cells Express Myodl and Myosin Heavy Chain

[0272] Consistent with the previous examples, no qualitative changes inPLA growth kinetics and morphology between the 8 patients used in thisstudy, suggesting that the isolated PLA populations are relativelyconsistent between all patients. PLA cells were isolated from rawlipoaspirates and induced using MM, containing hydrocortisone.Myogenesis by PLA cells was specific to the myogenic conditions used inthis study, as no differentiation was observed in non-inductive controlmedium, or in media inductive for alternate mesodermal lineages (i.e.osteogenic and adipogenic). Furthermore, no osteogenic or adipogenicdifferentiation was noted in PLA cells induced for up to 6 weeks in MM.

[0273] To confirm PLA myogenic potential, the expression of establishedmuscle-specific markers was determined by immunohistochemistry.Differentiation of myogenic precursors and stem cells into myogenicprecursor cells can be confirmed by the expression of severaltranscription factors, that include MyoD1, Myf-5, myogenin andstructural proteins such as myosin heavy chain (Butler-Browne, et al.,1990 Anat. Embryol. (Berl) 181:513-522; Thornell, et al., 1984 J.Neurol. Sci. 66:107-115; Megeney, et al., 1996 Genes Dev. 10:1173-1183;Seale and Rudnicki 2000 Dev. Biol. 218:115-124; Tapscott, et al., 1988Science 242:405-41 1; Weintraub, et al., 1991 Science 251:761-766;Molkentin and Olson 1996 Curr. Opin. Genet. Dev. 6:445-453). Commitmentto the myogenic lineage was identified by staining cells with amonoclonal antibody specific to MyoD1. Nuclear expression of MyoD1 inPLA cells was observed at 1, 3 and 6 weeks induction with MM, suggestinginitiation of the myogenic differentiation pathway in these cells (FIG.16, Panels A to C). Similar to the PLA results, nuclear expression ofMyoD1 was observed in positive control SKM cells as early as 1 weekpost-induction with MM and increased MyoD1 expression was observed inSKM cells by 6 weeks induction. In contrast to induced PLA cells, MyoD1expression was not observed in PLA cells treated for 1, 3 and 6 weekswith CM (FIG. 16, Panels D to F). Similarly, no MyoD1 expression wasobserved in HFF cells treated with MM . The expression of MyoD1 ininduced PLA cells suggests that these cells have initiated a program ofmyogenic differentiation.

[0274] To further confirm myogenesis, cells were stained with amonoclonal antibody specific to skeletal muscle myosin heavy chain(myosin), in order to identify terminally differentiated myoblasts(Butler-Browne, et al., 1990 Anat. Embryol. (Berl) 181:513-522;Thornell, et al., 1984 J. Neurol. Sci. 66:107-115). Consistent with thenuclear expression of MyoD1, PLA cells induced with MM also expressedmyosin (FIG. 17, Panels A to C). However, in contrast to MyoD1, myosinexpression was restricted to later induction time points (3 and 6 weeksonly), consistent with the expression of this marker in mature, fullydifferentiated myoblasts (Butler-Browne, et al., 1990 Anat. Embryol.(Berl) 181:513-522; Thomell, et al., 1984 J. Neurol. Sci. 66:107-115).Similar to our MyoD1 results, no myosin expression was observed in PLAcells cultured in CM (FIG. 17, Panels D to F) or in HFF cells inducedwith MM. Extensive myosin expression was also observed in SKM positivecontrols induced for 3 and 6 weeks with MM. Taken together, theexpression of both MyoD1 and myosin in induced PLA cells suggests thatthese cells possess myogenic potential.

[0275] Terminal differentiation of myogenic precursors is accompanied bythe fusion of the differentiated myoblast into long, multi-nucleatedmyotubes. Therefore, we examined induced PLA cultures for the formationof putative myotubes. Treatment of PLA cells with MM for a minimum ofthree weeks resulted in the formation of long multi-nucleated cells(FIG. 18A). The number and size of these multi-nucleated cells graduallyincreased with induction time with multi-nucleated cells observed in allPLA cultures at 6 weeks induction. No fusion was observed at 1-weekpost-induction with MM Furthermore, multi-nucleation was not observed inPLA cells cultured for similar time periods in CM or in HFF cellstreated with MM . To confirm the myogenic origin of these putativemyotubes, the expression of myosin was examined in PLA cultures at 6weeks post-induction. As shown in FIG. 18B, multi-nucleated PLA cells at6 weeks also expressed the myosin heavy chain. The formation ofmulti-nucleated cells expressing myosin upon induction with MM suggeststhat PLA cells underwent fusion to form myotubes and further confirmstheir myogenic potential in vitro.

RT-PCR Analysis

[0276] Finally, myogenic differentiation was confirmed using RT-PCR(FIG. 19). Consistent with our immunohistochemistry data, RT-PCRanalysis confirmed the expression of MyoD1 in PLA cells induced for 1, 3and 6 weeks in MM. In contrast, MyoD 1 expression was not observed inPLA cells cultured in CM nor in HFF cells induced with MM. Low levels ofmyosin expression were observed in induced PLA cells at 3 weeks, whileincreased expression of this marker was seen at 6 weeks post-inductionwith MM. Myosin was not detected in these cells after 1 week ofinduction and was supportive of the immunohistochemistry results. Theexpression of myosin was specific to induced PLA cells as no expressionwas detected in control PLA cells or in myo-induced HFF cells. TheRT-PCR results confirm our immunohistochemistry data and further supportthe myogenic potential of PLA cells.

Quantitation of Myogenic Differentiation by PLA Cells

[0277] In order to determine the degree of myogenic differentiation byinduced PLA cells, the immunohistochemistry data was quantitated. To doso, the number of MyoD1- or myosin-positive cells was counted as anindicator of myogenic marker expression level and expressed as apercentage of total PLA cells counted±the standard error of the mean (%total PLA±SEM). The number of MyoD1-positive PLA cells upon MM inductionis shown in FIG. 20. Low levels of MyoD1-positive PLA cells wereobserved after 1 week induction in MM (4.11±0.51% total PLA cells). By 3weeks post-induction, 10.11±3.85% of the total PLA cells were MyoD1positive while 15.37±4.33% of the total PLA cells were MyoD1 positive at6 week post-induction. Based on the cell count, a 2.4-fold increase inthe number of MyoD1 -positive cells was observed within the first 3weeks of induction. In contrast to the first 3 weeks, MyoD1 expressionlevels only increased 1.5-fold in the last 3 weeks of myogenicinduction. The greater number of MyoD1-positive cells in the first 3weeks of induction relative to the last 3 weeks may reflect the earlyrole this regulatory factor plays in myogenic differentiation (Megeney,et al., 1996 Genes Dev. 10:1173-1183; Seale and Rudnicki 2000 Dev. Biol.218:115-124; Tapscott, et al., 1988 Science 242:405-411; Weintraub, etal., 1991 Science 251:761-766).

[0278] In contrast to myo-induced PLA cells, no appreciable myogenicdifferentiation was observed at any time point upon treatment of PLAcells with CM or in HFF cells with MM, confirming the specificity of theinduction conditions. To confirm if the increase in MyoD1 expression inPLA cells over time was significant, statistical analysis was performedusing a one-way ANOVA. Comparison of the MyoD1 experimental values only,from 1 to 6 weeks, yielded statistical significance (P<0.001; F=18.9).In addition, analysis of 1 and 3 week MyoD1 levels using an unpairedt-test confirmed a significant difference (p=0.0021). A reduced level ofsignificance was determined between 3 and 6 weeks (p=0.0335) and islikely a reflection of the reduced role MyoD1 plays in maturingmyoblasts. Finally, the increased expression of MyoD1 in theexperimental group versus controls within each differentiation timeperiod was found to be statistically significant using a one-way ANOVA(p<0.0001).

[0279] A time-dependent increase in the number of myosin-positive PLAcells was also observed upon induction with MM (FIG. 21). Negligiblelevels of myosin expression were observed at 1 week post-induction,consistent with expression of this protein in maturing myoblasts.Following 3 weeks induction, 3.88±0.46% of the total PLA cells countedwere positive for myosin expression, while 8.42±0.71% were myosinpositive at 6 weeks, a 2.2-fold increase in the number ofmyosin-positive cells in the last 3 weeks of induction. The increasednumber of myosin expressing cells from 3 to 6 weeks post-induction wasgreater than that measured for MyoD1 and is consistent with a shift fromdifferentiating to maturing myogenic cells (Butler-Browne, et al., 1990Anat. Embryol. (Berl) 181:513-522; Thornell, et al., 1984 J. Neurol.Sci. 66:107-115). No myosin expression was observed in PLA cellscultured with CM or in HFF cells induced with MM, confirming thespecificity of the induction conditions. Analysis of the increase inmyosin expression levels from 1 to 6 weeks confirmed statisticalsignificance (one-way ANOVA—P<0.0001; F=75.5). Statistical significancewas also observed using an unpaired t-test to compare 3 and 6 weekmyosin expression levels only (p<0.0001). As in the MyoD1 studies,statistical analysis of both PLA and control cultures confirmedstatistical significance (one-way ANOVA—P<0.0001). Finally, myogenicdifferentiation levels, as measured using both MyoD1 and myosinexpression levels, were found to be consistent from patient to patient.Furthermore, regression analysis did not demonstrate a significantcorrelation of myogenic differentiation with patient age (MyoD1,correlation=0.27; myosin, correlation=0.30)

Discussion

[0280] Muscle loss due to trauma, vascular insult, tumor resection ordegenerative muscle disease such as muscular dystrophy represents asignificant clinical problem with few solutions. For focal muscle loss,vascularized muscle transplantation has been performed, but incumbentdonor site morbidity is both cosmetically and functionally limiting.System muscle disorders, such as degenerative muscle loss, are generallyconsidered to be fatal disorders resulting in progressive muscle loss,diaphragmatic paralysis or dysfunction and eventual suffocation. Currenttherapeutic approaches, such as gene therapy, have proven unsuccessfulthus far. However, recent developments in the field of tissueengineering may allow eventual replacement or repair of both focal andgeneralized muscle tissue loss.

[0281] Two cell types are generally considered candidate cells formuscle tissue engineering: embryonic stem cells and post-natally derivedprogenitor cells or stem cells. Unfortunately, ethical issues andpotential problems with cell regulation have limited the use ofembryonic stem cells (Baker, et al., 1996 Curr. Top. Dev. Biol.33:263-279; Dinsmore, et al., 1996 Cell Transplant 5:131-143; Lenoir, N.2000 Science 287:1425-1427; Rohwedel, et al., 1994 Dev. Biol.164:87-101; Young, FE 2000 Science 287:1424). Post-natal skeletal muscleprogenitors or satellite cells have been introduced for the treatment ofDuchenne's muscular dystrophy by Myoblast Transfer Therapy (MTT)(Karpati, et al., 1989 Am J. Pathol. 135:27-32; Law, et al., 1988 MuscleNerve 11:525-533; Rando, et al., 1995 Exp. Cell Res. 220:383-389;Partridge, et al., Nature 273:306-308). Although potentially beneficial,the practical use of satellite cells is limited primarily due to cellavailability (such cells must be harvested from viable donor muscletissue), as well as decreased self-renewal potential with increasing age(Rando, et al., 1994 J. Cell Biol. 125:1275-1287; Satoh, et al., 1993 J.Histochem. Cytochem. 41:1579-1582; Schultz and Lipton 1982 Mech. AgeingDev. 20:377-383). In addition to satellite cells, mesenchymal stem cellsderived from bone marrow (MSCs) have also been reported to have myogeniccapability under special culture conditions (Ferrari, et al., 1998Science 279:1528-1530; Wakitani, et al., 1995 Muscle Nerve18:1417-1426).

[0282] In this study, we show that human Processed Lipoaspirate (PLA)cells obtained from Suctioned-Assisted Lipectomy (SAL) have myogenicpotential in vitro. Immunohistochemical and RT-PCR analyses reveal thatPLA cells induced with MM express both MyoD1 and myosin heavy chain,markers that are expressed in skeletal muscle precursors undergoingdifferentiation and maturation. MyoD1 expression in PLA cells is highestduring the first 3 weeks of induction, consistent with its early role inmyogenic differentiation. A time-dependent increase in myosin is alsoobserved, with the highest number of myosin-positive cells observedduring the latter stages of differentiation (i.e. 3 to 6 weekspost-induction). Such an increase may reflect the maturation of PLAcells into myoblasts. Consistent with terminal differentiation andmyoblast fusion, long, multi-nucleated myotubes, expressing myosin, arefirst observed at three weeks post-induction, with the number and thesize of these multi-nucleated cells increasing over time. Finally,immunohistochemical quantification showed that approximately 15% of PLAcells undergo myogenesis.

[0283] In post-natal life, mature skeletal muscle fibers cannotregenerate if damaged. In response to muscle injury or in individualswith chronic degenerative myopathies, satellite cells, located betweenthe sarcolemma and the basal lamina of the muscle fiber, activate tobecome myogenic precursor cells. These precursors divide and fuse torepair the damaged muscle (Campion, DR 1984 Int. Rev. Cytol.87:225-251). However, the number of satellite cells within mature muscleis only 1-5% of the total cell number and their self-renewal potentialdecreases with age (Schultz and Lipton 1982 Mech. Ageing Dev.20:377-383; Alameddine, et al., 1989 Muscle Nerve 12:544-555). For focalmuscle loss, vascularized muscle transplantation has been performed, butincumbent donor site morbidity is both cosmetically and functionallylimiting. Furthermore, for systemic muscle diseases, autologous skeletaltissue transplantation cannot be used because of the generalized natureof the disease process. Therefore, other cell-based therapeuticapproaches are required.

[0284] One such emerging treatment strategy is Myoblast Transfer Therapyor MTT. Myoblast Transfer Therapy involves implanting large numbers ofhealthy myoblasts. This method was first performed in 1978 and has beenshown to be a promising treatment for Duchenne's muscular dystrophypatients (Karpati, et al., 1989 Am J. Pathol. 135:27-32; Law, et al.,1988 Muscle Nerve 11:525-533; Rando, et al., 1995 Exp. Cell Res.220:383-389; Partridge, et al., Nature 273:306-308). Althoughtheoretically beneficial for muscle tissue replacement or augmentation,its success has been limited (Rando, et al., 1994 J. Cell Biol.125:1275-1287; Satoh, et al., 1993 J. Histochem. Cytochem.41:1579-1582). As an alternative, multipotential stem cells have becomepromising candidates for future cell-based therapeutic strategies sincethey can rapidly proliferate in culture and retain the ability todifferentiate into several mesodermal cell types (Caplan 1991 J. Orthop.Res. 9:641-650; Pittenger, et al., 1999 Science 284:143-147).

[0285] Previous reports have demonstrated that mesodermal stem cells canbe isolated from both prenatal and post-natal organisms (Ferrari, etal., 1998 Science 279:1528-1530; Caplan 1991 J. Orthop. Res. 9:641-650;Elmer, et al., 1981 Teratology 24: 215-223; Swalla, et al., 1986 Dev.Biol. 116: 31-38; Hauschka, et al., 1974 Dev. Biol. 37: 345-68; Solursh,et al., 1981 Dev. Biol. 83: 9-19; Nakahara, et al., 1991 Exp. Cell Res.195: 492-503; Goshima, et al., 1991 Clin. Orthop. 274-283; Goshima, etal., 1991 Clin. Orthop. 298-311; Benayahu, et al., 1989 J. Cell Physiol.140: 1-7; Bennett, et al., 1991 J. Cell Sci. 99: 131-139; Calcutt, etal., 1993 Clin. Res. 41: 536A; Lucas, et al., 1992 In Vitro Cell Dev.Biol. 28: 154A; Lucas, et al., 1993 J. Cell Biochem. 17E: 122). Williamset al. has shown that post-natal cells isolated from skeletal muscletissue possess adipogenic, osteogenic, chondrogenic and myogenicpotential (Williams, et al., 1999 Am Surg. 65:22-26). Moreover, severalgroups have demonstrated the differentiation of Mesenchymal Stem Cells(MSCs) obtained from both human and animal bone marrow into adipogenic,osteogenic and chondrogenic lineage cells (Pittenger, et al., 1999Science 284:143-147; Grigoriadis, et al., 1988 J. Cell Biol.106:2139-2151; Beresford, et al., 1992 J. Cell Sci. 102:341-351; Cheng,et al., 1994 Endocrinology 134:277-286; Johnstone, et al., 1998 Exp.Cell Res. 238:265-272; Yoo, et al., 1998 J. Bone Joint Surg. Am.80:1745-1757). These findings suggest that bone marrow and skeletalmuscle may be a promising source of stem cells. However, there aredrawbacks to the use of bone marrow and skeletal muscle as sources ofmyogenic cells. Bone marrow procurement is painful and yields a lownumber of MSCs, often requiring ex vivo expansion prior to clinical use.Moreover, only a few stem cells can be obtained from skeletal musclewithout a functional loss to patients.

[0286] In this example we demonstrate the expression of establishedmyogenic markers by adipose-derived stem cells (MyoD1, myosin,multi-nucleation), confirming and quantitating their myogenic potential.Since adipose tissue is plentiful and liposuction procedures arerelatively safe with minimal patient discomfort, human adipose-derivedstem cells can provide an an additional source of multi-lineage cells,together with those obtained from bone marrow and skeletal muscle, fortreating muscular disorders.

[0287] While the expression of myogenic markers in stem cells was shown,the exact origin of these cells cannot be confirmed. It is possible,though unlikely, that our results are due to the contamination of theadipose compartment with satellite cells or myogenic precursors from anon-adipose tissue source. One possibility is the contamination of theadipose compartment with myogenic precursor cells from skeletal muscle.However, it is very unlikely from a technical standpoint that theinvesting fascia of the skeletal muscle could be entered with theblunt-tip liposuction cannula. Another possibility is the contaminationof the adipose compartment by MSCs from the peripheral blood.Conflicting reports have been presented as to the presence of MSCs inperipheral blood (Lazarus, et al., 1997 J. Hematother. 6:447-455; Huss2000 Stem Cell 18:1-9), although we believe that the level of myogenesisobserved in our study is inconsistent with the low percentage of MSCsthat might be contributed by peripheral blood. Finally, while no clearmarker exists for the identification of satellite cells and myogenicprecursors, MyoD1 is one of the earliest markers expressed duringdifferentiation and has been used to identify myogenic precursors(Weintraub, et al., 1991 Science 251:761-766; Grounds, et al., 1992 CellTiss. Res. 267:99-104; Sassoon, D A 1993 Develop. Biol. 156:11). Asshown in FIG. 16, MyoD1 expression was not observed in non-induced PLAcultures, suggesting that our results are not due to the presence ofmyogenic precursor cells in the PLA, but are due to the myogenicdifferentiation of a multi-lineage stem cell.

[0288] The goal of skeletal muscle tissue engineering is the treatmentof intrinsic skeletal muscle diseases and the loss of skeletal musclefollowing trauma or ischemia. Present medical and surgical therapies forthese disorders are either ineffective or impractical. The use of humanPLA cells in these areas is promising. Human PLA cells are plentiful,easily obtainable with minimal morbidity and discomfort and exhibitmyogenic potential. As such, these cells may have important applicationsfor myogenic tissue engineering and repair.

[0289] While the degree of myogenic differentiation of PLA cells isrelatively low compared to observed levels of adipogenic and osteogenicdifferentiation (Zuk, P., et al., 2001 Tissue Engineering 7:209-226),application of exogenous factors such as passive and active mechanicalforces (Periasamy, et al., 1989 Biochem. J. 257:691-698; Vandenburgh andKaufman 1981 J. Cell Physiol. 109:205-214; Vandenburgh 1983 J. CellPhysiol. 116:363-371; Vandenburgh, et al., 1988 In Vitro Cell Dev. Biol.24:166-174; Vandenburgh 1989 In Vitro Cell Dev. Biol. 25: 607-616) andrefinement of culture conditions may augment myogenic differentiation,making these cells clinically useful.

EXAMPLE 11

[0290] The following description provides adipose-derived stem cellswhich differentiate into osteogenic, chondrogenic, adipogenic, myogenic,and neurogenic tissues. The description also provides methods forisolating and inducing differentiation of said stem cells.

Materials and Methods

[0291] All materials were purchased from Sigma (St. Louis, Mo.), VWR(San Dimas, Calif.) and Fisher Scientific (Pittsburgh, Pa.) unlessotherwise stated. All tissue culture reagents were purchased from LifeTechnologies (New York, N.Y.). Fetal Bovine Serum (FBS) and Horse Serum(HS) were purchased from Hyclone (Logan, Utah) and Life Technologies,respectively. 1,25-dihydroxyvitamin D₃ was purchased from BioMol(Plymouth Meeting, Pa.).

Cell Lines

[0292] Normal human osteoblasts (NHOsts), normal human chondrocytes fromthe knee (NHCK) and a population of MSCs from human bone marrow werepurchased from Clonetics (Walkersville, Md.). The murine 3T3-L1preadipocyte cell line (Green and Meuth 1974 Cell 3:127-133) wasobtained from ATCC (Rockville, Md.). The human neuroendocrine cell line,PC 12, was the generous gift of Dr. Harvey Herschman (UCLA, Los Angeles,Calif.).

Antibodies

[0293] A monoclonal antibody to human osteocalcin was purchased fromTaKaRa Shizo Co. (Japan). The polyclonal antibodies to human osteopontin(αOP-LF123), osteonectin (αON—LF37), biglycan, (αBG—LF51), decorin(αDEC—LF136) and alkaline phosphatase (αAP) were obtained from Dr. LarryFisher (NIH). Monoclonal antibodies to MAP2 (αMAP), neurofilament 70(αNF70) and τ-tau (αtau) were purchased from Leinco Technologies (St.Louis, Mo.). Monoclonal antibodies to trk-a (αTRK) and NeuN (αNeu) werepurchased from Santa Cruz Biotech (Santa Cruz, Calif.) and Chemicon(Temecula, Calif.), respectively. Polyclonal antibodies to glialfibrillary acidic protein (αGFAP) and were purchased from Dako andStressgen (Victoria, BC), respectively. Secondary antibodies conjugatedto alkaline phosphatase were obtained from Zymed, while secondaryantibodies conjugated to FITC were purchased from BioSource (CamarilloCalif.).

Cell Harvest, Culture and Differentiation Conditions

[0294] Adipose-derived stem cells (PLA) cells were obtained from rawlipoaspirates and cultured as described in a previous study (Zuk, 2001Tissue Engineering 7(2):209-226). Adipose-derived stem cells and 3T3-L1cells were maintained in non-inductive Control medium (Table 5). NHOst,MSC and NHCK cells were maintained in specialized commercial Controlmedia (Clonetics). Adipose-derived stem cells cells were induced towardthe desired mesenchymal lineages as outlined in Table 5. MSCs wereinduced using commercial control medium supplemented with the growthfactors outlined in Table 5. 3T3-L1 cells were induced toward usingAdipogenic Medium (AM). NHOst and NHCK cells were induced usingcommercially available induction media (Clonetics).

Histology, Immunohistochemistry and Indirect Immunofluorescence

[0295] Indirect Immunofluorescence (IF): PLA cells and MSCs wereprocessed for IF as described in Zuk, P. et al., 2001 Tissue Engineering7:209-226 using monoclonal antibodies to specific CD markers (Table 6).

[0296] Histology and Immunohistochemistry (IH): To confirmlineage-specific differentiation, differentiated cells were processed asdescribed in Zuk, P. et al., 2001 Tissue Engineering 7:209-226, usingthe following histological assays: Alkaline Phosphatase (osteogenesis),Oil Red O (adipogenic) and Alcian Blue (chondrogenic). In addition, PLAnodules induced toward the chondrogenic lineage were processed by IH forthe expression of collagen type 2, keratan sulfate (KS) andchondroitin-4-sulfate (CS), as previously described in Zuk, P. et al.,2001 Tissue Engineering 7:209-226.

Spectrophotometric Assays

[0297] Alkaline Phosphatase (AP): Triplicate samples of PLA cells weredifferentiated in Osteogenic Medium (OM) for up to 6 weeks. Cells werewashed with PBS and harvested into PBS/0.1% Triton X-100 (PBS/TX10O). APenzyme activity was assayed using a commercial AP enzyme kit (Sigma) andmeasured at an absorbance of 405nm. Total protein in each sample wasmeasured based on the Bradford method (Bradford 1976 Anal. Biochem.72:248-254) using a BCA Protein Assay Kit (Pierce, Rockford, Ill.). APactivity was expressed a nmol p-nitrophenol produced/minute/ug protein.The assay was calibrated using standard p-nitrophenol solutions.Differentiated MSC and NHOst cells were assayed as positive controlswhile non-induced PLA cells were assayed as a negative control. Valuesare expressed as the mean±SD.

[0298] Total Calcium (Ca²⁺): Triplicate samples of PLA cells weredifferentiated in OM as described above. Cells were washed with PBS (noCa2+, no Mg²⁺) and harvested in 0.1N HCl. Cells were extracted for aminimum of 4 hours at 4° C. and centrifuged at 1000×g for 5 minutes.Total calcium in the supernatant was determined using Sigma kit #587 andmeasured at A575 nm. The assay was calibrated using calcium standardsolutions. Total protein was determined and the samples were expressedas mM Ca/ug protein.

[0299] Differentiated MSC and NHOst cells were assayed as positivecontrols, while non-induced PLA cells were assayed as a negativecontrol. Values are expressed as the mean±SD.

[0300] Glycerol-3-Phosphate Dehydrogenase (GPDH): Triplicate samples ofPLA cells were differentiated in AM for up to 5 weeks. The cells wereharvested in 25 mM Tris-Cl, 1 mM EDTA (pH 7.5), 0.1 mM β-mercaptoethanoland sonicated for 5 sec and 40 W to lyze. The suspension was centrifugedat 10000×g and GPDH in the supernatant assayed by measuring theoxidation of NADH at A340 nm, according to the method of Wise and Green(Wise, 1979). One unit of GPDH was defined as the oxidation of 1 nmol ofNADH per minute. Samples were normalized with respect to protein andexpressed as units GPDH/ug. Differentiated MSC and 3T3-L1 cells wereassayed as positive controls while non-induced PLA cells were assayed asa negative control. Values are expressed as the mean±SD.

[0301] Dimethyldimethylene Blue (DMMB): Triplicate samples of PLA cellswere differentiated in Chondrogenic Medium (CM) for up to 3 weeks usingestablished micromass protocols (Ahrens, et al., 1977 Develop. Biol.60:69-82; Denker, et al., 1995 Differentiation 59:25-34; Reddi, et al.,1982 Prog. Clin. Biol. Res. 110(Part B):261-268). PLA nodules wereharvested and assayed for the sulfated glycosaminoglycans keratansulfate (KS) and chondroitin sulfate (CS) according to the method ofFarndale et al.(Farndate, et al., 1986 Biochimica et Biophysica Acta883:173-177). The assay was calibrated by the use of standard KS and CSsolutions. Samples were normalized with respect to protein and expressedas μg KS or CS per μg protein. Non-induced PLA cells were assayed as anegative control. Values are expressed as the mean±SD.

RT-PCR Analysis

[0302] PLA cells were induced toward the five lineages outlined in Table5 for defmed time periods. Total cellular RNA was isolated from thedifferentiated cells using a commercially available kit (QiaEasy,Qiagen). RNA was reverse transcribed using an oligo-dT primer andMMLV-Reverse Transcriptase (Promega, Madison, Wis.) for 60 minutes at42° C. PCR amplification was performed by the addition of Taq buffer(Promega), 2.5 mM MgCl₂, 1 mM dNTPs and 50 pmol of the appropriateprimer set (Table 7). The mix was incubated for 1 minute at 94° C. and2.5 units of Taq polymerase (Promega) was added. PCR was performed for40 cycles (1 minute—94° C., 1 minute—57° C., 1 minute—72° C.: finalextension—5 minutes at 72° C.). All primer sequences were determinedusing established GenBank sequences and the Primer3 program. PCRreactions with primers designed to the housekeeping gene β-actin wereamplified for 35 cycles as an internal control. The sequence of each PCRproduct was confirmed using automated sequencing. Non-induced PLA cellswere examined as a negative control. Lineage-specific cell lines wereanalyzed as a positive controls for the osteogenic, adipogenic andchondrogenic lineages. Total human skeletal muscle and brain RNA werereverse-transcribed and amplified by PCR as a positive control for themyogenic and neurogenic lineages, respectively.

Western Blotting

[0303] PLA cells were differentiated and harvested in 1% SDS. Lysateswere homogenized and total protein assayed. Equivalent amounts ofprotein from each lineage were denatured for 5 minutes at 100° C. in SDSLoad Buffer (0.5M Tris-Cl (pH 6.8), 1%SDS, 1 mM DTT, 50% Glycerol, 1%Bromophenol Blue). Lysates were resolved by polyacrylamide gelelectrophoresis (10% separating gel, 5% stacking gel), according tostandard protocols. Proteins were transferred overnight tonitrocellulose membranes and the membranes blocked for a minimum of 60minutes in Western Blocking Buffer (WBB: 5% non-fat milk, 1×PBS, 0.1%Tween-20). Membranes were incubated for a minimum of 60 minutes in WBB,supplemented with the following antibodies: osteogenesis: αOP, αON,αCNI, αDEC and αBG and adipogenesis: αG4 and αLEP. Membranes were alsoincubated with antibodies to the transferrin receptor and the solubleheat shock protein HSC70 as internal controls. Membranes were washed aminimum of 3 times with PBS/0.1% Tween-20 and then incubated for 60minutes with WBB supplemented with the appropriate secondary antibodyconjugated to alkaline phosphatase. The membranes were washed, asdescribed above, and the secondary antibodies visualized using acommercial kit (CSPD Ready-To-Use, Tropix, Bedford, Mass.) according tothe manufacturer. Non-induced PLA cells were also analyzed as a negativecontrol.

Neurogenic Differentiation

[0304] Immunohistochemistry: Subconfluent PLA cells were cultured inPreinduction Medium (DMEM, 20% FBS, 1 mM β-mercaptoethanol) for 24hours. Following preinduction, cells were induced for up to 8 hours inNeurogenic Medium (NM) and assessed by IH in order to determine specificneurogenic lineages (Table 8).

[0305] RT-PCR: PLA cells were pre-induced for 24 hours in PreinductionMedium, followed by induction in NM for up to 9 hours. PLA samples wereharvested, RNA isolated (QiaEasy, Qiagen) and analyzed by RT-PCR for theexpression of specific neurogenic genes (Table 7) as detailed above.

Isolation and Analysis of PLA Clones

[0306] Clone Isolation: PLA cells were plated at extremely lowconfluency in order to result in isolated single cells. Cultures weremaintained in Control medium until proliferation of single PLA cellsresulted in the formation of well-defined colonies. The single PLA-cellderived colonies were harvested using sterile cloning rings and 0.25%trypsin/EDTA. The harvested clones were amplified in Cloning Medium (15%FBS, 1% antibiotic/antimycotic in F12/DMEM (1:1)).

[0307] Confirmation of Multi-lineage capacity: Expanded clones wereanalyzed for multi-lineage potential as described earlier (see Histologyand Immunofluorescence).

[0308] Molecular Characterization: Clones were analyzed by RT-PCR forthe expression of several lineage-specific genes as described above.

Results Stem Cells Share Many Similarities With MSCs

[0309] In order to characterize the PLA population further, cells wereexamined using indirect IF and compared to a commercial population ofhuman MSCs. MSCs have been shown to express a unique set of cell surfacemarkers that can be used to help identify this stem cell population(Table 6) (Bruder, et al., 1998 J. Orthop. Res. 16:155-162; Cheng, etal., 1994 Endo 134:277-286; Jaiswal, et al., 1997 J. Cell Biochem.64:295-312; Pittenger, et al., 1999 Science 284:143-147). Like MSCs, PLAcells expressed several of these proteins (FIG. 23), supporting thecharacterization of these cells as stem cells. Approximately 100% of thePLA and MSC cultures were positive for the expression of CD29, CD44,CD90 and CD105/SH2 with high expression levels for each of these markersbeing observed in both cell populations. Both cell populations alsoexpressed the SH3 antigen, which, together with SH2, is considered aspecific marker for MSCs (Haynesworth, et al., 1992 Bone 13:69-80). Inaddition, the majority of PLA cells and MSCs were also positive for thetransferrin receptor, CD71, indicating that a fraction of these cellpopulations were replicating. PLA and MSCs did not express thehaematopoietic lineage markers, CD31 and CD34. A small number of PLAsamples did show negligible staining for CD45, although the number ofCD45-positive cells did not exceed 5% of the total PLA cell number.Unlike MSCs, no staining for the adhesion molecule CD58 was observed inPLA cells. Flow cytometric analysis for CD marker expression confirmedthe IF results (FIG. 24). Taken together, the immunofluorescent and flowresults demonstrate several similarities in CD expression profilesbetween PLA populations and bone marrow-derived MSCs.

PLA Cells Undergo Distinct Changes Upon Osteogenic Induction

[0310] In this Example, we demonstrate that PLA cells undergo distinctproliferative, synthetic and mineralization phases upon osteogenicinduction. In order to characterize the osteogenic capacity of PLA cellsfurther, the proliferation of osteo-induced PLA cells was measured andcorrelated to AP activity and calcium phosphate formation (FIG. 25,Panel A). PLA cell number increased upon initiation of osteogenicdifferentiation (day 1 to day 3), however, negligible AP and Von Kossastaining was observed (FIG. 25, Panel B). A linear increase in PLA cellnumber was observed from day 3 to day 9 and minimal AP staining wasobserved up until day 13. PLA proliferation rates leveled off brieflybetween day 13 and day 15, a phenomenon that was observed in several PLApopulations. A dramatic increase in AP activity was seen between day 15and day 19 and the first appearance of calcium phosphate deposits wereobserved by 3 weeks induction. An enhanced rate of PLA proliferation wasmeasured from day 15 to day 25 and coincided with a time-dependentincrease in both AP and VK staining. In addition, the formation ofmultilayered nodular structures and increased matrix synthesis were alsoobserved during this time period . PLA cell number decreased from day 25onward and was accompanied by the development of intemodular regionslacking adherent cells, together with increased matrix mineralization.Together, these results suggest that PLA cells may undergo distinctproliferative and metabolic phases as osteogenic differentiationproceeds.

Alkaline Phosphatase Activity and Time-Dependent Increase in MatrixMineralization

[0311] Bone formation in vivo is a complex process involving morphogens,hormones and growth factors. Recent work has questioned the efficacy ofsynthetic glucocorticoids, like dexamethasone (Dex), in mediatingosteogenesis. Glucocorticoids appear to inhibit the action of severalosteogenic genes including osteocalcin, CBFA-1 and CNI (as reviewed inCooper, et al., 1999 J. Endocrinol. 163:159-164). It is well establishedthat bone tissue and osteoprogenitor cells are targets of vitamin Daction (Chen, et al., 1983 J. Biol. Chem. 258:4350; Chen, et al., 1979J. Biol. Chem. 254:7491; Narbaitz, et al., 1983 Calcif. Tiss. Int.35:177) and this metabolite stimulates both AP activity and CNIsynthesis by human bone cell populations (Beresford, et al., 1986 Endo119:1776-1785). Therefore, PLA cells were induced using two osteogenicmedia compositions: containing either dexamethasone (Dex at 10⁻⁷ M) or1,25-dihydroxyvitamin D₃ (VD at 10⁻⁸ M). AP activity and Ca²⁺accumulation were measured over time using commercial kits andnormalized with respect to protein and/or time.

[0312] Induced PLA, MSC and NHOst cells were measured for AP activityand stage-specific induction levels presented in Table 9. AP activity inPLA cells, resulting from either Dex or VD-induction, first appeared at3 weeks and undifferentiated PLA cells exhibited negligible AP levels atall time points (FIG. 26, Panel A). AP activity from 3 to 6 weeks wasbi-phasic upon both Dex and VD stimulation of PLA cells, with peakactivities at days 21 and 42 and decreasing levels at day 35. VDinduction of PLA cells resulted in higher enzyme activities at 3, 4 and5 weeks, while no significant difference could be measured between thetwo induction conditions at 6 weeks. Maximum AP levels were detected at3 weeks in VD-induced PLA cells, whereas no significant maximum wasdetected upon Dex treatment. Moreover, PLA cells appeared to be moreresponsive to VD stimulation at 3 weeks with an enhanced level of enzymeinduction being measured in these cells compared to Dex treatment(17.2-fold induction/Dex vs. 71.3-fold induction/VD). Like PLA cells,negligible AP activity was measured in Dex-treated MSCs until day 21. Incontrast to PLA cells, Dex stimulation of MSCs resulted in higher enzymeactivities at 3, 4 and 5 weeks. The overall pattern of MSC AP activityobserved under Dex and VD induction was similar to VD-induced PLA cells(i.e. bi-phasic). However, decreasing levels were measured at day 28 inMSCs rather than day 35. Moreover, maximum enzyme activities in MSCswere detected at a later differentiation stage (i.e. 5/6 weeks).Finally, as observed in PLA cells, induction of MSCs from 2 to 3 weeksresulted in the greatest induction of AP activity. However,dexamethasone, rather then VD treatment, affected enzyme levels more inthese cells (54.2-fold induction/Dex vs. 1.1-fold induction/VD). Thepattern of AP enzyme activity was dramatically different in NHOstosteoblasts. Maximum AP levels were observed at 7 days in these cellsand enzyme levels decreased after this time point to reach minimumlevels at 6 weeks. Negligible enzyme activity could be detected inVD-treated osteoblasts at day 35 and 42. Furthermore, no significantdifference in AP activity could be measured from day 7 to day 28 undereither induction condition.

[0313] Induction of PLA cells and MSCs with either Dex or VD resulted ina time-dependent increase in matrix mineralization (FIG. 26 and Table10). Consistent with AP activity, PLA cells were more responsive toVD-induction, producing a greater overall increase in matrixmineralization (122-fold/VD vs. 56-fold/Dex), with maximum levelsdetected at 6 weeks. A similar effect, was also observed in VD-treatedMSCs, although maximum levels were reached one week earlier. As with APactivity, negligible mineralization in both PLA cells and MSCs wasobserved until 3 weeks. A true effect of induction condition was onlyobserved in PLA cells at 6 weeks, with VD-treated cells associated withsignificantly more calcium phosphate. In contrast to PLA cells,induction condition significantly affected mineralization in MSC sampleswith Dex treatment resulting in greater calcium levels early indifferentiation (3 and 4 weeks), consistent with the AP levels underthis induction condition. This trend was reversed at 5 and 6 weeks, withVD resulting in enhanced mineral levels. Interestingly, a decrease inCa²⁺ was observed in both Dex- and VD-treated MSCs at 28 days andappeared to correlate with the decrease in AP activity at this timepoint. In contrast to PLA cells and MSCs, maximum Ca²⁺ accumulationoccurred at 2 weeks in induced NHOst cells and decreased beyond thistime point, consistent with observed NHOst AP activity. Like MSCs, Dexinduction resulted in greater Ca²⁺ levels at all induction points withthe exception of 5 weeks. Control osteoblasts were associated withminimal levels of Ca²⁺, indicating that these cells do not spontaneouslymineralize without appropriate induction. Taken together, the AP andCa²⁺ spectrophotometric data further supports the osteogenic phenotypeof PLA cells.

Osteo-Induced PLA Cells Express Osteocalcin and CBFA-1

[0314] To confirm the osteogenic phenotype of PLA cells at the molecularlevel, osteo-induced PLA cells were analyzed using RT-PCR and Westernblotting. For RT-PCR analysis, PLA cells were induced for increasingtime periods in OM containing either Dex or VD. Dex and VD-induced MSCswere also analyzed, in addition to NHOst cells. Osteogenicdifferentiation did not appear to affect PLA cells, as β-actin levelsdid not differ significantly from control cells (FIG. 27). Inductionwith Dex or VD did not significantly affect the expression of themajority of the genes examined. However, a dramatic effect was observedin the expression of the bone-specific gene, OC. OC expression was notdetected in Dex-treated and control PLA cells, nor in control andDex-induced NHOst osteoblasts. Treatment of PLA cells with VD produced abi-phasic OC expression pattern. Negligible levels of OC were detectedat 4 and 14 days of induction, whereas a significant increase wasobserved at day 7. Finally, a relatively consistent level of OCexpression was detected from day 21 to day 42. Elimination of Dex andreplacement with VD for the last 48 hours of PLA induction wassufficient to overcome the effects of Dex and induce significant levelsof OC expression. In contrast to the RT-PCR results, analysis using agene microarray detected a slight increase in OC expression inDex-treated PLA cells versus non-induced controls (FIG. 27, Panel B). OCwas not specific to osteo-induced MSCs, as detectable levels wereobserved in control MSCs. Dex treatment of MSCs appeared to increase OClevels at 4 and 7 days and, like control cells, was followed by adecrease at 2 and 4 weeks. Unlike PLA cells, VD induction of MSCs. didnot result in a bi-phasic OC expression pattern. Rather, expressionlevels of this gene appeared to remain consist across the 4 weekinduction period and were elevated when compared to Dex treatment.

[0315] In addition to OC, expression of the bone-specific transcriptionfactor CBFA1 was observed in osteo-induced PLA cells using RT-PCR. CBFA1was expressed at all induction points and no discernible effect onexpression was observed upon Dex or VD induction. In addition, CBFA1expression was not specific to osteo-induced PLA cells as a lower levelof this gene was seen in controls. However, an increased level of CBFA1expression (approx. two-fold) was measured in osteo-induced PLA cellsusing gene arrays (FIG. 27, Panel B). Like PLA cells, CBFA1 wasexpressed in Dex- and VD-induced MSCs at each induction point, inaddition to being expressed in undifferentiated MSC controls at adecreased level. CBFA1 expression was more restricted in osteoblasts,detected in 4 week osteo-induced NHOst cells only. AP expression wasobserved at all time points in differentiated and control PLA cells,MSCs and NHOst cells. In addition to CBFA1 and AP, high levels of CNIwere observed in these cells. While, no appreciable difference in CNIexpression level was seen upon Dex or VD induction of PLA cells byRT-PCR, gene array analysis confirmed a decrease in CNI level uponosteogenic induction (FIG. 27, Panel B). In addition to OC, CBFA1, APand CNI, differentiated PLA cells, MSCs and NHOsts expressed othermarkers consistent with bone differentiation, including OP and ON . Asseen with CNI, decreased levels of ON and OP were also measured inDex-treated PLA cells using arrays (FIG. 27, Panel B). An increasedexpression of the transcription factor, PPARγ1 was also observed inDex-treated PLA and MSCs when compared to non-induced controls. Inaddition, a lower level of PPARγ1 was seen in the early stages of VDinduction of PLA cells (day 4 to day 14) and was followed by increasedexpression beyond four weeks induction. Osteogenic induction did notresult in the expression of genes consistent with fat and cartilagedifferentiation (PPARγ2 and CNII, respectively). Together, theexpression of bone-specific OC and other genes characteristic ofosteogenic differentiation in osteo-induced PLA cells further supportsthe osteogenic capacity of these cells.

[0316] Finally, osteogenic differentiation by PLA cells was confirmed atthe protein level by immunofluorescent analysis and Western blotting.Osteo-induced PLA cells (OM/Dex) were analyzed by IF for the expressionof OP, ON and OC. MSCs, induced under identical conditions were alsoexamined as a control. As shown in FIG. 28, non-induced andosteo-induced PLA cells specifically expressed both OP and ON (FIG. 28,Panel A). OP was distributed evenly throughout control and inducedcells, in addition to a distinct perinuclear concentration. Noextracellular OP staining could be observed. A punctate intracellularpattern was also observed for ON in both cell types. In addition,increased nuclear staining for this protein was also observed incontrols. Upon osteo-induction, ON staining appeared to increase inareas of concentrated cells and was expressed both intracellularly andextracellularly. No expression of OC could be observed in non-inducedcells and was consistent with the RT-PCR results. A small percentage ofthe osteogenic PLA cells appeared to express low levels of OCintracellularly. Similar expression patterns for these proteins wereobserved in MSCs. Control and induced MSCs expressed high levels of bothOP and ON. Like PLA cells, a punctate intracellular and nuclear stainingpattern were observed for ON with the nuclear staining decreasing uponinduction (FIG. 28, Panel B). Control and induced MSCs expressed OPintracellularly. However, unlike PLA cells, no perinuclear concentrationfor this protein could be seen. Finally, consistent with the RT-PCRdata, both control and induced MSCs expressed OC.

[0317] To confirm the expression of osteogenic proteins by Westernanalysis, PLA cells were maintained in OM for 7, 14 and 21 days andlyzed. Cell lysates were analyzed for CNI, OP, ON, Decorin and Biglycanexpression. In addition, lysates were also analyzed for the transferrinreceptor (TfR) as internal controls. OC was not assessed due to thesmall size of the protein (6 kDa). Osteogenic differentiation did notappear to alter the expression of the TfR as equivalent levels were seenin osteo-induced cells and controls maintained for 3 weeks in Controlmedium. Comparable levels of CNI, Decorin and Biglycan were seen inosteo-induced PLA cells at all three induction periods. In addition,these proteins were also seen in controls, suggesting that bothdifferentiated and undifferentiated PLA cells are associated with aproteinaceous ECM. Like these matrix proteins, both ON and OP were seenin differentiated cells and undifferentiated controls. However, ONlevels appeared to decrease upon initial induction of PLA cells andreturned to control levels by 3 weeks. In addition, osteogenic inductionwas accompanied by a slight increase in OP at day 21. Taken together,the immunofluorescent and Western data confirms the expression ofproteins consistent with osteogenic differentiation by PLA cells.

PLA Cells Undergo Adipogenic Differentiation

[0318] Adipogenic differentiation is associated with the growth arrestof preadipocytes before commitment to the differentiation program(reviewed in Ailhaud, et al., 1992 Annu. Rev. Nutr. 12:207-233;MacDougald and Lane 1995 Annu. Rev. Biochem. 64:345-373; Smyth, et al.,1993 J. Cell Sci. 106:1-9). To determine the correlation between PLAproliferation and adipogenic differentiation, PLA cells were inducedtoward the adipogenic lineage in AM for up to 3 weeks and cell numbersdetermined, together with the degree of differentiation using Oil Red Ostaining. The differentiation (i.e. appearance of intracellular lipidvacuoles) first appeared as early as 4 days induction. Consistent withthe commitment of preadipocytes, no appreciable increase in PLA cellnumber was detected over the course of adipogenic induction (FIG. 29,Panel A) despite a time-dependent increase in Oil Red O staining/lipidaccumulation levels (FIG. 29, Panel B). Differentiation levels weregreatest in culture regions in which the PLA cells were confluent and incontact with one another. These results suggest that the commitment ofPLA cells to the adipogenic lineage is influenced by cell-cell contactand coincides with growth arrest.

[0319] Adipose conversion of preadipocyte cells lines, such as 3T3-L1,is also characterized by an increase in the activity of lipogenicenzymes, including glycerol-3-phosphate dehydrogenase (GPDH) (Wise, etal., 1979 J. Biol. Chem. 254:273-275). PLA cells were therefore inducedwith AM and the level of GPDH activity determined. 3T3-L1 cells weresimilarly induced as a positive control. Initial induction of PLA and3T3-L1 cells (day 4 to day 7) resulted in comparable GPDH activities andwere similar to control PLA levels (FIG. 30). Induction of PLA cells fortwo weeks resulted in a decrease in enzyme activity and no significantdifference between control, induced PLA and 3T3-L1 cells was observed.This decrease was also observed in adipo-induced MSCs. Adipogenicdifferentiation beyond 2 weeks resulted in an increase in GPDH activityspecifically in induced PLA and 3T3-L1 cells. Enzyme activity leveledoff between 4 and 5 weeks and was significantly higher than control PLAcells. The time-dependent increase in GPDH activity correlated with theincreased percentage of lipid-filled PLA cells within adipo-inducedcultures (FIG. 29) and was consistent with adipogenic differentiation bythese cells.

[0320] To confirm PLA adipogenesis, adipo-induced cells were analyzed byRT-PCR. As shown in (FIG. 31), PLA induction resulted in the expressionof the adipose-specific transcription factor PPARγ2. Expression ofPPARγ2 was observed at day 7 and the levels appeared to remainconsistent throughout the remainder of the 5 week induction period. Noexpression of this gene was detected in non-induced PLA cells. Inaddition to PPARγ2, low levels of the adipogenic genes LPL and aP2 wereexpressed in induced PLA cells. Low levels of these genes were observedupon early adipose induction (4 days) and were followed by significantincrease at 1 week. Increased levels were maintained in these cells asfar as 5 weeks induction. Basal expression of LPL and aP2 was alsoobserved in control PLA cells, although at a significantly lower levelthan induced samples. Like osteo-induced PLA cells, PPARγ1 was expressedin adipo-induced cells. However, the expression pattern of this geneappeared to be distinct from osteo-induced cells, with low expressionlevels observed at early time periods (day 4 to 14) followed byincreased expression from 3 to 6 weeks. Adipogenic induction of MSCsresulted in similar gene expression patterns. Like PLA cells, PPARγ2expression was specific to adipo-induced MSCs and did not appear at theearliest stages of induction. Extremely low levels of aP2 and LPL werealso observed in control MSCs and adipogenic induction resulted in asignificant increase in these genes beyond 7 days. However, in contrastto PLA cells, PPARγ1 was not observed in control MSCs. Rather,expression of this transcription factor was restricted to adipo-MSCs.Furthermore, the expression pattern of this gene paralleled that ofPPARγ2, with no expression being observed until 1 week of induction.Finally, expression of these genes was examined in 3T3-L1 cells inducedtoward the adipogenic lineage or induced via growth to confluence.Expression of aP2 and LPL were observed in adipo-induced 3T3-L1 cells,while an apparent inhibition of PPARγ2 expression was seen. Adipogenicdifferentiation of PLA cells, MSCs and 3T3-L1 cells did not result inthe expression of the bone-specific gene, OC and the cartilagenousmarker, CNII, confirming the specificity of the adipogenic inductionconditions. In summary, the restricted expression of PPARγ2 byadipo-induced PLA cells, together with the increased expression of aP2and LPL upon induction supports the in vitro adipogenic capacity ofthese cells.

Chondrogenic Differentiation

[0321] Induction of PLA cells cultured under micro-mass conditions withCM resulted in the formation of well-defined, compact nodules consistentwith those seen upon chondrogenic induction of MSCs (Johnstone, et al.,1998 Exp. Cell Res. 238:265-272; Yoo, et al., 1998 J. Bone Joint Surg.Am. 80:1745-1757) Chondrogenic differentiation of PLA cells wasdependent upon cell density and induction conditions. Specifically, PLAnodules formed in induction medium containing TGFβ1 alone, while theaddition of dexamethasone increased the size of TGFP1-induced PLAnodules. Nodule formation was not observed in the presence ofdexamethasone alone. Attempts to initiate PLA chondrogenesis inmonolayer culture was unsuccessful. To assess the ECM produced bychondrogenic PLA cells, nodules were examined by IH for the expressionof CNII and sulfated proteoglycans. PLA nodules, induced for 14 days inCM, stained positively using Alcian Blue, which specifically identifiessulfated proteoglycans (FIG. 32, Panel A, AB). In support of this, 14day PLA nodules also stained positively using monoclonal antibodiesspecific for keratan and chondroitin-4-sulfate (Panel A, KS and CS,respectively). Expression of CNII was also observed in these nodules.Alcian Blue and CNII staining were also detected in sections of humancartilage and were not seen in high density PLA cultures maintained inControl medium, confirming the specificity of our histologic and IHprotocols.

[0322] In addition to IH staining for KS and CS, the level of thesesulfated proteoglycans was measured using a quantitativedimethyldimethylene blue assay (FIG. 32, Panel B). PLA nodules and NHCKcontrols were predigested with papain to eliminate possible interferenceby proteins and glycoproteins prior to assay. A time-dependent increasein KS and CS was observed in PLA nodules up to 2 weeks of chondrogenicinduction. A slight decrease was observed at 3 weeks for both PGs.Non-induced PLA cells, maintained under high-density conditions, werealso associated with an ECM containing these proteoglycans. Furthermore,control PLA cells at 4 and 7 days induction contained more KS and CS incomparison to induced samples. However, significantly more proteoglycanaccumulation was observed in induced PLA cells at days 14 and 21.

[0323] Treatment of PLA cells for 2 weeks in CM resulted in theexpression of several genes consistent with chondrogenesis as shown byRT-PCR (FIG. 33). CNII expression was observed specifically in inducedPLA cells and was restricted to day 7 and 10. RT-PCR analysis confirmedthe presence of both the IIA and IIB splice variants of CNII, althoughthe IIB variant only is shown in FIG. 33. A low level of CNII expressionwas also observed upon chondrogenic induction of NHCK controls. Asimilar expression pattern to CNII was observed in induced PLA nodulesusing primers designed to the amino terminus of the large proteoglycan,aggrecan (AG). Expression of this proteoglycan was also observed usingprimers to the carboxy terminus (PG). However, aggrecan expression bythe PLA nodule using the carboxy primers was observed from day 7 to day14. In support of the PLA results, chondrogenic induction of NHCKcontrols also resulted in the expression of aggrecan using both primersets. Finally, like CNII, aggrecan expression was specific to inducedPLA nodules and NHCK cells. In addition to CNII, chondrogenic inductionof PLA nodules resulted in the specific expression of CNX, a marker ofhypertrophic chondrocytes, at day 14 only. In contrast to this, noexpression of CNX could be observed in NHCK controls and may be due totheir derivation from articular cartilage. PLA cells were alsoassociated with additional collagen types. Both induced and control PLAcells expressed CNI and CNIII. While the majority of PLA samplesexamined exhibited a restricted collagen expression pattern (day 4only), a few PLA samples showed expression of CNI and CNIII up to day14. Induced PLA cells also expressed the proteoglycans, decorin andbiglycan and the gene Cbfa-1. Expression of these genes was observedthroughout the entire induction period and was also seen in control PLAcells. While decorin and biglycan levels remained consistent, a slightdecrease in CBFA-1 levels appeared at later stages of induction (i.e.days 10 and 14). No expression of OC was seen at any time point,confirming the absence of osteogenic differentiation. Taken together,the specific expression of CNII, aggrecan and CNX in induced PLAnodules, in addition to the presence of keratan- and chondroitin-sulfatewithin the ECM supports the chondrogenic phenotype of these cells.

PLA Cells Express Myod1, Myf5, Myogenin And Myosin Transcipts

[0324] MSCs from rat have been shown to possess myogenic potential(Saito, 1995; Wakitani, 1995). To examine if PLA cells possess thiscapacity, cells were examined for the expression of the early myogenicregulatory factors, myoD1 and myf5, in addition to myogenin and themyosin heavy chain, a later marker of myogenic differentiation.Expression of myod1, myogenin and myosin was observed at all inductionpoints, while expression of myf5 appeared to be restricted to 1 and 3weeks only (FIG. 34). Consistent with the role of myodl myogenicdetermination, increased levels of this gene were observed at 1 week.Furthermore, while myogenin levels appeared to remain consistent, a timedependent increase in expression was detected for myosin, consistentwith the expression of this protein in mature myoblasts. In support ofthe PLA results, expression of these four myogenic genes was alsoobserved in samples of total RNA prepared from human skeletal muscle.Therefore the expression of these myogeic regulatory proteins indicatespossible myogenic differentiation by PLA cells.

Clones Derived from Single PLA Cells Possess Multi-Lineage Capacity:Adipose Derived Stem Cells (ADSCs)

[0325] The presence of multiple mesodermal potential in PLA cells isstrong support for the characterization of these cells as stem cells.However, this phenomenon may simply be due to the contamination bylineage-specific precursors. To determine if this is the case, PLA cellswere cultured at a low enough confluence to promote the formation ofcolonies derived from single PLA cells. Several multi-lineage cloneswere isolated and those possessing tri-lineage potential were termedtermed Adipose Derived Stem Cells or ADSCs. Like PLA cells, ADSCs werefibroblastic in morphology. Following expansion, no evidence of othercell morphologies could be observed, confirming the homogeneity of ADSCcultures. Analysis of 500 PLA clonal isolates confirmed differentiationpotential in approximately 6% of the total number of clones examined.Seven ADSC isolates exhibited tri-lineage potential, differentiatinginto cells of the osteogenic, adipogenic and chondrogenic lineages(Table 11). In addition to tri-lineage ADSCs, several dual-lineageclones (O/A, O/C and A/O) and single adipogenic lineage clones were alsoisolated (FIG. 35). A qualitative increase in differentiation level, asmeasured by histologic staining, was observed in all PLA clonalpopulations . Finally, isolation and expansion of tri-lineage ADSCs didnot alter the CD expression profile as shown by IF, nor coulddifferences be detected in the dual lineage clones (FIG. 36). RT-PCRanalysis of tri-lineage ADSCs confirmed their multi-lineage potential(FIG. 36).

[0326] Induction of ADSCs in OM/VD for 2 to 4 weeks resulted in theexpression of OC and 3 and 4 weeks only, consistent with osteo-inducedPLA cells, in which no OC expression could be detected at 2 weeks. Inaddition to OC, expression of ON, OP, CNI and AP were seen at allinduction points. Like PLA cells, expression of OC was specific toinduced ADSCs, nor could the fat marker PPARγ2 be detected in bothinduced and control clones. Fat induction of ADSCs for 2 and 4 weeksresulted in the specific expression of aP2 and LPL. Interestingly, adramatic decrease in PPARγ2 was observed in fat ADSCs, expressed weaklyat 4 weeks only. As seen in the heterogenous PLA population, noosteogenic differentiation was detected in adipogenic ADSCs. Finally,expression of aggrecan, CNX, decorin and biglycan was detected upon 2weeks of chondrogenic induction. No expression of CNII could be observedin these cells at this induction point. Like PLA cells, expression ofaggrecan and CNX was restricted to chondrogenic ADSCs, nor could OCexpression be detected. Together with the IH data, the RT-PCR resultsconfirms the multi-lineage capacity of ADSC isolates and suggests thatthe multi-lineage capacity of the PLA population may be due to thepresence of a putative stem cell population.

PLA Cells May Possess Neurogenic Potential

[0327] The mesodermal embryonic layer gives rise to several connectivetissues while the overlying ectoderm is the progenitor of multipleneural tissues and cell types. Recent evidence suggests that MSCs can beinduced toward non-mesodermal lineages, differentiating to cells withputative neurogenic potential (Deng, et al., 2001 Biochem. Biophys. Res.Commun. 282:148-152; Sanches-Ramos, et al., 2000 Exp. Neurol.164:247-256; Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370). It ispossible that the similarities between PLA and MSCs may extend beyondmesodermal potential. Therefore, PLA cells were induced toward theneuroectodermal lineage based on the protocol of Woodbury et al.(Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370) and examined forthe expression of neural markers, including NSE, trk-a and MAP-2, or theexpression of GFAP and GaIC, markers of astrocytes and oligodendrocytes,respectively. To induce the PLA cells, subconfluent cultures werepre-treated with 1 mM β-mercaptoethanol (βME) and 20% FBS for a maximumof 24 hours (pre-induction), followed by induction in serum-free mediumwith 5-10 mM βME (Neurogenic Medium/NM) for up to 8 hours. Pre-inductiondid not change the fibroblastic morphology of the PLA cells (FIG. 38,Panel A—PLA/0 hrs). A morphologic change was noted as early as 30minutes induction in NM, with 10% of the cultures assuming aneuronal-like phenotype. No morphological changes were observed if FBSwas added to the NM. Sixty minutes of induction increased the proportionof neuronal-like cells to 20% of the culture. Induction for three hoursincreased this phenotype to a maximum of 70% and no significant increasewas observed beyond this induction time. NM-induced PLA cells underwentretraction, forming compact cells bodies with multiple extensions. Cellbodies became more spherical and cell processes exhibited secondarybranches with increasing induction time (Panel A—PLA/2 hrs vs. PLA/8hrs). Induction in NM resulted in significant expression of NSE, trk-aand NeuN, consistent with the neuronal lineage (Panel B). Virtually 100%of the PLA culture stained positively for both NSE and trk-a. Incontrast to the NSE results, not all PLA cells appeared to be NeuNpositive, and may represent a more defined subset of neuronal-like cellswithin the PLA culture. No expression of the mature neuronal markersMAP-2 and NF-70 were observed, suggesting that induced PLA cellsrepresent an early developmental stage. In addition, induced PLA cellsdid not express GalC and GFAP, indicating that PLA cells did notdifferentiate into oligodendrocytes and astrocytes, respectively.Finally, control PLA cells did not express any neuronal,oligodendrictyic or astrocytic markers, confirming the specificity ofour induction conditions and staining protocol.

[0328] To further assess the resulting lineage upon NM induction, PLAcells were analyzed by RT-PCR (FIG. 38, Panel C). PLA cells induced for4.5 hours in NM expressed significant amounts of nestin, an intermediatefilament protein expressed in significant quantities in neural stemcells and precursors (Lendahl, 1990). Nestin expression was alsodetected in non-induced PLA cells and in total RNA prepared from humanbrain. No expression of ChaT, a marker of peripheral nerves, wasobserved in NM-induced cells or in brain. In addition, NM-induced PLAcells did not express GAD65, a marker of mature neurons and wasconsistent with the lack of IH staining using antibodies to thisneuronal stage (eg. MAP-2, NF-70). As seen in the IH results, PLA cellsalso did not express GFAP. Similar expression patterns were observed inPLA cells induced for 9 hours. The expression of nestin, NSE, NeuN andtrk-a, together with the lack of ChaT, or GFAP expression suggests thatPLA cells may be capable of differentiating into an early neuronalphenotype, characteristic of the CNS. Thus the PLA cultured in NM candifferentiate into an ectodermal lineage. Furthermore, recent data byLumelsky et al. (Lumelsky, N., et al. 2001 Science 292:1389) show thatan embryonic stem cell can be induced to differentiate into a cell thatexpresses nestin. The nestin-postive cell was further characterized tobe a pancreatic precursor cell. Therefore, Lumelsky's data suggests thatnestin-positive cells can differentiate into both an endodermal lineageand an ectodermal lineage. An endodermal phenotype can be furtherconfirmed by the additional expression of one or more of the following:insulin, glucose transporter 2, islet amyloid polypeptide, GATA4, GATA6,albumin, tyrosin aminotransferase. TABLE 5 Lineage-specificdifferentiation induced by media supplementation Medium Media SerumSupplementation Control DMEM 10% FBS none Adipogenic DMEM 10% FBS 0.5 mMisobutyl-methylxanthine (AM) (IBMX), 1 μM dexamethasone, 10 μM insulin,200 μM indomethacin, 1% antibiotic/ antimycotic Osteogenic DMEM 10% FBS0.1 μM dexamethasone, 50 μM (OM) ascorbate-2-phosphate, 10 mMβ-glycerophosphate, 1% antibiotic/ antimycotic Chondrogenic DMEM 1% FBS6.25 μg/ml insulin, 10 ng/ml (CM) TGFβ1, 50 nM ascorbate-2- phosphate,1% antibiotic/ antimycotic Myogenic DMEM 10% FBS, 0.1 μM dexamethasone,50 μM (MM)  5% HS hydrocortisone, 1% antibiotic/ antimycotic NeurogenicDMEM none 5-10 mM β-mercaptoethanol (NM)

[0329] TABLE 6 Monoclonal antibodies to CD antigens: Reported cellspecificity and distribution CD Antigen Clone Cell Specificity 29Integrin β1 MAR4 broad distribution—lymphocytes, monocytes, granulocytesNOT on erythrocytes 31 PECAM-1 9G11 endothelial cells, platelets,monocytes, granulocytes, haematopoietic precursors 34 — 581 endothelialcells, some tissue fibroblasts, haematopoietic precursors 44 Pgp-1G44-26 leucocytes, erythrocytes, epithelial cells, platelets 45 LCA HI30leucocytes, haematopoietic cells 58 LFA-3 L306.4 widedistribution—haematopoietic cells, endothelial cells, fibroblasts 71 TfRH68.4 most dividing cells 90 Thy-1 5E10 immature CD34+ cells, cellscapable of long term culture, primitive progenitor cells 105  Endoglin —endothelial cells, B cell precursors, MSCs SH3 — — mesenchymal stemcells

[0330] TABLE 7 Oliogonucleotide primer sequences and expected PCRproduct sizes Product Lineage Gene Oligonucleotide primers size BONEOsteonectin (ON) 5′ TGTGGGAGCTAATCCTGTCC 400 bp 3′ TCAGGACGTTCTTGAGCCAGTOsteopontin (OP) 5′ GCTCTAGAATGAGAATTGCACTG 270 bp3′ GTCAATGGAGTCCTGGCTGT Osteocalcin (OC) 5′ GCTCTAGAATGGCCCTCACACTC 300bp 3′ GCGATATCCTAGACCGGGCCGTAG Bone sialoprotein (BSP)5′ GCTCTAGAATGAAGACTGCTTTAATT 185 bp 3′ ACTGCCCTGAACTGGAAATC Corebinding factor 5′ CTCACTACCACACCTACCTG 320 bp α-1 (CBFA-1)3′ TCAATATGGTCGCCAAACAGATTC Collagen I (CNI) 5′ GAGAGAGAGGCTTCCCTGGT 300bp (α1 chain) 3′ CACCACGATCACCACTCTTG Alkaline phosphatase5′ TGAAATATGCCCTGGAGC 475 bp (AP) 3′ TCACGTTGTTCCTGTTTAG FAT aP25′ TGGTTGATTTTCCATCCCAT 150 bp 3′ TACTGGGCCAGGAATTTGAT LPL5′ GAGATTTCTCTGTATGGCACC 275 bp 3′ CTGCAAATGAGACACTTTCTC PPAR gamma15′ GCTCTAGAATGACCATGGTTGAC 3′ ATAAGGTGGAGATGCAGGCTC PPAR gamma25′ GCTGATATGGGTGMACTCTG 3′ ATAAGGTGGAGATGCAGGTTC PPAR delta5′ GCCAACGGCAGTGGCAATGTC 3′ TTAGTACATGTCCTTGTAGATCTC CARTILAGE CollagenII (α1′ chain) 5′ ATGATTCGCCTCGGGGCTCC 260 bp 3′ TCCCAGGTTCTCCATCTCTGAggrecan 5′ GCAGAGACGCATCTAGAAATT 505 bp 3′ GGTAATTGCAGGGAACATCATDecorin 5′ CCTTTGGTGAAGTTGGAACG 300 bp 3′ AAGATGTAATTCCGTAAGGG Biglycan5′ TGCAGAACAACGACATCTCC 475 bp 3′ AGCTTGGAGTAGCGAAGCAG Collagen X5′ TGGAGTGGGAAAAAGAGGTG 600 bp 3′ GTCCTCCAACTCCAGGATCA MUSCLE MyoD15′ AAGCGCCATCTCTTGAGGTA 500 bp 3′ GCGCCTTTATTTTGATCACC Myf55′ CCACCTCCAACTGCTCTGAT 250 bp 3′ GGAGTTCGAGGCTGTGAATC Myogenin5′ TGGGCGTGTAAGGTGTGTAA 130 bp 3′ TTGAGCAGGGTGCTTCTCTT Myosin5′ TGTGAATGCCAAATGTGCTT 750 bp 3′ GTGGAGCTGGGTATCCTTTGA NERVE CHaT5′ TACAGGCTCCACCGAAGACT 375 bp 3′ AGCAGAACATCTCCGTGGTT Synaptophysin(SYN) 5′ TTCAGGCTGCACCAAGTGTA 350 bp 3′ CAGGGTCTCTCAGCTCCTTG GlialFibrillary Acidic Protein 5′ AATGCTGGCTTCAAGGAGAC 405 bp (GFAP)3′ CCAGCGACTCAATCTTCCTC GAD65 5′ TGGCGATGGGATATTTTCTC 300 bp3′ GCACTCACGAGGAAAGGAAC Nestin 5′ GGAGTCGTTTCAGATGTGGG 240 bp3′ AGCTCTTCAGCCAGGTTGTC

[0331] TABLE 8 Assessment of neurogenic differentiation by PLA cells:antibodies and established neurogenic lineages Antibody Name ProteinLineage NeuN Neuron-specific nuclear Neurons & Neural proteinprogenitors NF-70 Neurofilament 70 kDa Neurons trk-A trk-A (NGFreceptor) Neurons MAP2 microtubule associated Neuron (mature) protein-2GalC galactocerebroside Oligodendrocytes GFAP glial acidic fibrillaryAstrocytes protein τ-tau tau Neurons, Oligodendrocytes, Astrocytes

[0332] TABLE 9 Alkaline phosphatase induction levels AP Induction(“x”-fold) Cell line day 14-21 day 21-28 day 28-35 day 35-42 PLA-Dex+17.2 +1.6 −2.9 +3.2 PLA-VD +71.3 −1.3 −1.9 +1.9 MSC-Dex +54.2 −1.2 +2.7NS MSC-VD NS −1.5 +3.5 +1.5 NHOst-Dex −1.4 NS −2.8 −1.2 NHOst-VD +1.4−2.2 −25.5 ND

[0333] TABLE 10 Quantitation of calcium phosphate levels Change inOverall Calcium Content Cell line (“x”-fold increase/decrease) PLA-Dex56 PLA-VD 122  MSC-Dex 12 MSC-VD 67 NHOst-Dex ND NHOst-VD ND

[0334] TABLE 11 Summary of Lineage-Specific ADSC Differentiation LineageSpecific Differentiation O, A, C O, C A, O A, C O only A only C only #ADSC 7 10 3 3 0 6 0 Clones

[0335] TABLE 12 Flow cytometric analysis of CD marker expression oncontrol PLA cells CD Antigen Geometric Mean  CD4 2.44  CD8 2.31  CD11c2.49 CD13 148.88 CD14 2.43 CD16 2.38 CD19 2.92 CD31 2.22 CD33 2.61 CD343.55 CD44 16.92 CD45 2.52  CD49d 5.33 CD56 2.66 CD61 3.68   CD62E 2.30CD71 3.76 CD90 25.96 CD104  2.31 CD105  8.39 CD106  2.45 SH3 8.95 STRO-131.26 −ve 2.59

Discussion

[0336] To further confirm if PLA cells represent a mesenchymal stem cellpopulation, we conducted an extensive molecular and biochemicalcharacterization of this cell population and several PLA clones termedAdipose-Derived Stem Cells, or ADSCs. PLA populations were inducedtoward multiple mesodermal lineages, including bone and fat, and theexpression of lineage-specific genes and proteins confirmed by RT-PCR,indirect immunofluorescence (IF) and Western blotting. In addition,established biochemical assays were used to measure the activities ofalkaline phosphatase, a marker for bone metabolism, the lipogenic enzymeglycerol-3-phosphate dehydrogenase (GPDH), together with theaccumulation of sulfated proteoglycans upon chondrogenic induction.Histological analysis and RT-PCR were also used to confirm themulti-lineage differentiation of ADSCs. Finally, the potential of PLAcells to differentiate into cells of the neurogenic lineage was alsoexamined.

[0337] We have demonstrated the multi-lineage capacity of theheterogenous PLA cell population and its clonal derivatives, ADSCs,obtained from human lipoaspirates. In agreement with this work, weconfirm that PLA cells and ADSC clones are capable of osteogenic,adipogenic, chondrogenic and myogenic differentiation as shown by theexpression of several lineage-specific genes and proteins. In additionto mesodermal lineages, PLA cells also appeared to undergodifferentiation to a lineage consistent with the neurogenic phenotype.Taken together, the molecular and biochemical data suggest that PLAcells may represent a putative stem cell population that can be isolatedfrom human adipose tissue.

PLA Cells Express a Similar Complement of CD Markers as Observed in MSCs

[0338] Characterization of a cell population can be accomplished throughidentification of unique proteins expressed on the cell surface. Severalgroups have subsequently characterized MSCs based on their expression ofcell-specific proteins (e.g. STRO-1, SH2, SH3, SH4) and “clusterdesignation” (CD) markers (Bruder, et al., 1998 J. Orthop. Res.16:155-162; Conget, et al., 1999 3. Cell Physiol. 181:67-73; Pittenger,et al., 1999 Science 284:143-147). This study confirms that a uniquecombination of cell surface proteins is expressed on PLA cells.Moreover, both PLA and MSC populations show similar expression profiles.Like MSCs, PLA cells expressed CD29, CD44, CD71, CD90, CD105/SH2, SH3and STRO-1 as shown by IF, in addition to CD13 as confirmed by FC (FIG.24). Like MSCs, PLA cells did not express CDs 4, 8, 11, 14, 16, 19, 31,33, 34, 45, 56, and 62E on the cell surface (FIG. 24). The similar CDprofiles suggest that PLA cells may be a stem cell population like MSCs.However, the degree of similarity may indicate that PLA cells are simplyan MSC population located within or contaminating the adiposecompartment. Lipoplasty results in the rupture of multiple blood vesselsand while vasoconstrictors are used to minimize blood loss, theprocessed PLA pellet may be MSCs obtained from the peripheral bloodsupply (Zvaifler, et al., 2000 Arthritis Res. 2:477-488). However, thereappear to be a few subtle distinctions between PLA and MSC populations.In contrast to MSCs, no expression of CD58 could be detected on PLAcells using IF, while expression was seen on MSCs (FIG. 23).Furthermore, MSCs have also been reported to express CD104, CD106 andCD140a (Bruder, et al., 1998 J. Orthop. Res. 16:155-162; Conget, et al.,1999 J. Cell Physiol. 181:67-73; Pittenger, et al., 1999 Science284:143-147). No expression of these CD antigens were detected on PLAcells using IF or FC (Table 12). These differences may indicate that thePLA population is a distinct population of stem cells. However, thepossibility that PLA cells are a clonal variant of MSCs cannot be ruledout.

PLA Cells Undergo Osteogenesis

[0339] The mesengenic process involves: 1) proliferation of progenitorcells, 2) commitment of these cells via the action of specific growthfactors and cytokines, 3) lineage progression into transitory cell typesexpressing specific genes and 4) terminal differentiation characterizedby the cessation of proliferation and biosynthesis of tissue-specificproducts (Bruder, et al., 1997 J. Cell Biochem. 64:278-294; Caplan 1994Clin. Plas. Surg. 21:429-435; Jaiswal, et al., 1997 J. Cell Biochem.64:295-312). Osteogenesis follows this pattern closely with osteogenicprecursors developing into mitotic pre-osteoblasts and secretoryosteoblasts, which lose their mitotic potential and form the matureosteocyte (Owen, et al., 1990 J. Cell Physiol. 143:420-430; Stein, etal., 1989 Conncet. Tissue. Res. 20:3-13). Therefore, osteogenicdifferentiation is characterized by distinct phases of proliferation,matrix synthesis/maturation and mineralization (Owen, et al., 1990 J.Cell Physiol. 143:420-430). Consistent with this, distinct phases wereobserved upon osteogenic differentiation of PLA cells. A relativelylinear growth rate was measured within the first week of induction, aperiod characterized by negligible AP activity and Ca²⁺ deposition.Proliferation rates increased between day 9 and day 13 and wereaccompanied by the appearance of AP by day 13. Proliferation ceasedtemporarily between day 13 and day 15 and no significant increase in APstaining was observed during this time point . An increase in cellnumber and enhanced AP staining was observed beyond 2 weeks induction.These findings are similar to the sequence of events in reportedcalvarial cultures in which cells first proliferate and then showelevated levels of AP (Aronow, et al., J. Cell. Physiol. 143:213-221;Owen, et al., 1990 J. Cell Physiol. 143:420-430). Moreover,glucocorticoids have been postulated to stimulate the proliferation ofosteogenic progenitors (Shalhoub, et al., 1989 Biochem. 28:5318-5322;Tenenbaum, et al., 1985 Endo 117:2211-2217. In addition to AP activity,significant levels of calcium were seen by 3 weeks and marked the onsetof the mineralization phase in PLA cells. Increased matrixmineralization was accompanied by a dramatic increase in AP staining andwas consistent with results found in rat calvarial cultures (Collin etal., 1992 Calcif. Tiss. Int. 50:175-183; Shalhoub, et al., 1989 Biochem.28:5318-5322). Increased mineralization was also accompanied by thecessation of proliferation (day 25), followed by a reduction in PLA cellnumber. This reduction was likely due to the increase in mineraldeposition and coincided with the increased appearance of ECM and theformation of cell-free intemodular zones . In support of this, ECMformation has been suggested to contribute to the shutdown ofproliferation by rat osteoblasts (Owen, et al., 1990 J. Cell Physiol.143:420-430) and rat marrow stromal cells (Malaval, et al., 1994 J.Cell. Physiol. 158:555-572). Taken together, the results suggest thatPLA cells possess distinct proliferative, synthetic and mineralizationphases during osteogenic differentiation.

[0340] Glucocorticoid excess and/or prolonged treatment in vivo isassociated with decreased bone formation (Baylink 1983 N. Engl. J. Med.309:306-308), possibly through a reduction of progenitor conversion toosteoblasts (Chyun, et al., 1984 Endo. 114:477-480). In contrast todexamethasone, treatment with vitamin D metabolites restores bonemineralization and bone formation by bone-derived cells in vitro(Beresford, et al., 1986 Endo. 119:1776-1785; Kanis, et al., 1982 inEndocrinology of Calcium Metabolism, ed. J A Parsons, New York: RavenPress, pp 321). Therefore, the effects of dexamethasone and1,25-dihydroxyvitamin D3 (VD) on PLA osteogenesis were examined. Thebone/kidney/liver isoform of AP catalyzes the cleavage of inorganic andorganic phosphates at alkaline pH. While its precise function during invivo osteogenesis is unclear, AP expression levels in pre-osteoblastsand MSCs are upregulated upon the onset of osteogenic differentiationand this enzyme thought to play a key role in matrix mineralizationthrough its pyrophosphatase activity (McComb, et al., 1979 AlkalinePhosphatase, New York: Plenum Press; Robison 1923 Biochem. J.17:286-293; Siffert 1951 J. Exp. Med. 93:415-426). Therefore, analysisof AP levels and matrix mineralization are important indicators ofosteogenesis. Based on this, these parameters were measured in inducedPLA samples and compared to similarly treated MSCs and human osteoblastsas controls.

[0341] While, the overall effect of osteogenic differentiation on APactivity and matrix mineralization appeared to be similar in PLA cellsand MSCs, the kinetics of enzyme activity and the response to inductionconditions differed depending on differentiation stage, suggesting thatthese two populations may possess distinct phenotypes. AP activityappeared in both PLA and MSC populations between 2 and 3 weeksinduction. VD treatment of PLA cells resulted in a significantly higherlevel of AP activity at 3 weeks versus Dex induction and a greater levelof enzyme induction from 2 and 3 weeks (17.2 fold/Dex vs. 71.3 fold/VD).This VD effect was seen at each differentiation stage. In contrast, theeffect of induction condition was reversed in MSCs, with Dex producinggreater AP activities at each differentiation stage. In addition todifferences in measured enzyme activity and induction level, thekinetics of AP activity differed between PLA cells and MSCs. AP activityin both Dex and VD-induced PLA cells was bi-phasic. Specifically, peakAP levels were measured under both induction conditions at 3 and 6 weeksand a decreased level detected at 5 weeks. Like PLA cells, a bi-phasicresponse was also observed in Dex-treated MSCs. However, the kinetics ofAP activity appeared to be accelerated in Dex-treated MSCs with peaksdetected at 3 and 5 weeks and a decrease in enzyme at 4 weeks. Moreover,a distinct bi-phasic pattern was not observed upon VD stimulation ofMSCs, lending further support to a putative distinction between thesetwo cell populations.

[0342] The reason for the biphasic response in PLA and MSC populationsis unclear. Time course studies using rat calvarial models have shownthat AP activity peaks early, during the deposition of the bony ECM, andis subsequently downregulated (Owen, et al., 1990 J. Cell Physiol.143:420-430; Rodan and Rodan 1984 in Bone and Mineral Research, ed. W.A., Amsterdam: Elsevier Science Publishers, pp. 244-285; Stein, et al.,1990 FASEB J. 4:3111-3123). A similar pattern is observed in marrowstromal cell cultures and correlates with advanced matrix mineralizationand terminal osteogenic differentiation into osteocytes (Bruder andCaplan 1990 Bone 11:189-198; Jaiswal, et al., 1997 J. Cell Biochem.64:295-312; Malaval, et al., 1994 J. Cell. Physiol. 158:555-572). Thedrop in PLA AP activity observed from 4 to 5 weeks correlates toincreasing calcium phosphate levels within the matrix. However, we knowof no studies in which AP levels are quantitated beyond this matrixsynthesis phase. Therefore, this study may be the first to examine APactivity in stem cells over an extended time period. It is possible thatthe pattern of PLA and MSC AP activity represents a stage-specificresponse to osteogenic induction. In support of this, increases in APhave been observed in VD-treated immature osteosarcoma cultures(Majeska, et al., 1982 J. Biol. Chem. 257:3362) whereas a dose-dependentinhibition was detected in more mature cells, an effect thought torepresent the return of a cell fraction to the osteoprogenitor pool ortheir differentiation to osteocytes, a cell population with low APactivity. Therefore, the decrease in AP levels in PLA and MSC samplesmay be due to the terminal differentiation of a cell fraction whereasthe second AP peak could be due to the delayed development of a fractionof osteogenic progenitor cells.

[0343] Consistent with the AP results, induction of PLA cells with VDproduced a greater overall increase in calcium levels compared todexamethasone. Like AP activity subtle distinctions in calciumaccumulation could be observed between PLA cells and MSCs. In support ofthe AP data, matrix mineralization by PLA cells was not observed until 3weeks induction. Beyond this time point, Dex stimulation did not appearto significantly affect the rate of matrix mineralization. However, adramatic increase was detected in VD-treated PLA samples, with 6 weeksamples containing significantly more calcium phosphate. This increasedmineralization rate occurred despite the fact that AP did not differdramatically between 3 and 6 week VD samples. Moreover, the decrease inAP activity observed between 4 and 5 weeks in PLA cells did nottranslate into decreases in calcium level. Rather, mineral accumulationcontinued to increase in these cells. This pattern has previously beenobserved in human MSCs (Jaiswal, et al., 1997 J. Cell Biochem.64:295-312). Like PLA cells, a time dependent increase in mineralizationwas observed in MSCs with a greater overall increase observed inVD-treated samples. The pattern of matrix mineralization in these cellscorrelated well with AP activity within the first 4 weeks of induction.Specifically, higher AP levels in Dex-treated MSCs resulted in greatercalcium accumulation. However, between 4 and 5 weeks induction adramatic shift takes place, with small increases in AP activity inVD-treated MSCs producing dramatic increases in calcium level. Moreover,AP activity in VD-induced MSCs were significantly lower than Dex-treatedcells, yet VD treatment resulted in dramatically more calcium,suggesting that MSCs became more sensitive to VD induction over time.Taken together, the appearance of AP upon osteogenic induction and theaccumulation of a mineralized ECM support the osteogenic phenotype ofPLA cells. In addition, differences observed in the kinetics and patternof these two markers indicates that the PLA population may be distinctfrom MSCs.

[0344] During osteogenesis, osteoblasts synthesize a wide repertoire ofno proteins that are incorporated into a surrounding ECM scaffold. Thecomposition of the matrix, together with the kinetics of secretion, helpdefine the unique properties of bone tissue and can be used to confirmosteogenic differentiation. However, with few exceptions, the actualmatrix proteins are not unique to bone. One of these exceptions is theprotein osteocalcin (OC). A highly conserved protein containing threeγ-carboxyglutamic acid residues, OC is an inhibitor of hydroxyapatiteformation in vitro, suggesting that this protein participates inmineralization (Boskey, et al., 1985 Calc. Tiss. Int. 37:75; Price, etal., 1976 Proc. Natl. Acad. Sci. USA 73:1447-1451). In support of this,OC is expressed by mature osteoblasts and its expression level risesdramatically during the mineralization phase (Collin, et al., 1992Calcif. Tiss. Int. 50:175-183; Malaval, et al., 1994 J. Cell. Physiol.158:555-572; Owen, et al., 1990 J. Cell Physiol. 143:420-430; Shalhoub,et al., 1992 J. Cell. Biochem. 50:425-440; Stein, et al., 1990 FASEB J.4:3111-3123). While OC is considered a relatively late marker ofosteoblast differentiation, it is expressed early in bone formation inmarrow stromal cell cultures before large amounts of matrix aresynthesized (Malaval, et al., 1994 J. Cell. Physiol. 158:555-572).Consistent with osteogenic differentiation, osteo-induced PLA cellsexpressed OC. However, its expression was dependent upon the compositionof the osteoinductive medium. Specifically, osteocalcin expression wasnot observed in non-induced PLA cells nor in PLA cells induced with OMcontaining dexamethasone. The lack of OC expression in Dex-treated PLAcells may be due to an inhibitory effect associated with glucocorticoids(Cooper, et al., 1999 J. Endocrinol. 163:159-164). In support of this,negligible levels of OC have been observed in rat MSCs and human bonecell cultures induced with dexamethasone (Beresford, et al., 1986 Endo.119:1776-1785; Leboy, et al., 1991 J. Cell Physiol. 146:370-378).Furthermore, OC was not observed upon induction of a human osteoblastcell line, NHOst, in this study (FIG. 27). In contrast to dexamethasoneinduction, OC expression was seen only upon VD stimulation and isconsistent with studies confirming VD-dependent increases in OCexpression by osteosarcoma cells (Price, et al., 1980 J. Biol. Chem.225:11660-11663) and its stimulation of the OC promoter (Lian, et al.,1988 Clin. Orthop. Rel. Res. 226:276-291; Yoon, et al., 1988 Biochem.27:8521-8526). In addition to its appearance upon VD induction, adistinct bi-phasic expression pattern of OC was observed. Consistentwith bone marrow MSCs, the appearance of OC was associated with aninitial stage of differentiation, appearing as early as 4 daysinduction. A dramatic increase in OC level was detected after one weekinduction. Induction for 2 weeks resulted in an apparent inhibition ofOC expression and was followed by increased expression beyond threeweeks. The reappearance of OC at three weeks was coincident with thesynthesis and mineralization of the surrounding ECM and may besupportive of the proposed role for OC in matrix calcification. Withregards to OC's biphasic pattern, a similar effect to that observed inAP expression may be occuring: i.e. a developmental stage-specificresponse to VD. In addition to VD, several other induction agents alsoexert stage-specific effects on osteogenesis, including TGFβ (Breen, etal., 1994 J. Cell. Biochem. 160:323-335). Similar to VD-induced PLAcells, OC was also detected in MSCs with several differences observed inOC expression pattern observed in these cells. First, in contrast to PLAcultures, a low level of OC expression was observed in non-induced MSCs.The basal level of OC expression in control MSCs was extremely low andis consistent with reports of constitutive OC expression in cultures ofrat MSCs (Malaval, et al., 1994 J. Cell. Physiol. 158:555-572). Second,OC expression was observed in Dex-treated MSCs. Finally, whileVD-induction increased the expression of OC in MSCs, no apparentbiphasic pattern was observed. Taken together, the expression ofbone-specific OC by osteo-induced PLA cells supports their osteogeniccapacity. In addition, the distinct pattern of OC expression and thedifferential response to induction factors observed between MSCs and PLAcells further suggests that these two populations may possess uniquephenotypes.

[0345] In addition to OC, osteo-induced PLA cells also expressed severalother genes characteristic of the osteogenic lineage, including OP, ON,CBFA1, AP and CNI. Cbfa-1 (core binding factor-1 or Osf-2) is atranscriptional regulatory factor encoded by the gene, CBFA1, a memberof the runt domain gene family (Kania, et al., 1990 Genes Dev.4:1701-1713). Isolated from the nuclear extracts of primary osteoblasts,the Cbfa1 factor has been shown to bind to the promoters of severalosteogenic genes, including OC, OP BSP and CN type I, thus acting as amaster regulator of osteoblast differentiation (Ducy, et al., 1997 Cell89:747-754). Moreover, mutations to the C-terminal region of human CBFA1is associated with Cleidocranial dysplasia (CCD), an autosomal-dominantcondition characterized by deformities in skeletal patterning (Jones, etal., 1997 Smith's Recongizable Patterns of Human Malformation, 5^(th)edition, Philadelphia: W B Saunders Company; Mondlos, et al., 1997 Cell89:773-779; Otto, et al., 1997 Cell 89:765-771; . Consistent with itsproposed role, both Dex and VD-induced PLA cells expressed CBFA1 at allinduction points and no significant difference in expression level wasobserved between the two induction conditions. In support of the PLAresults, CBFA1 was also expressed in osteo-induced MSCs and wasrestricted to a late differentiation stage in NHOst cells. Finally, bothundifferentiated PLA cells and MSCs expressed low levels of this growthfactor. However, osteogenic induction of PLA cells resulted in anapproximate 2-fold increase in CBFA1 expression as confirmed using genearrays. Moreover, recent studies in developing mice have suggested thatCbfa1 is expressed in progenitors of both the osteogenic andchondrogenic lineages (Ducy, et al., 1997 Cell 89:747-754). Therefore,the expression of CBFA1 in control PLA cells may represent basal geneexpression in cells with a progenitor phenotype.

[0346] Like CBFA1, the expression of OP, ON, AP and CNI was observed incontrol and osteo-induced PLA cells, MSCs and NHOsts throughoutdifferentiation. Expression of CNI in these cell types appeared to beequivalent under each induction condition using RT-PCR. However,decreased expression of this gene was detected in osteo-induced PLAcells using gene arrays and is consistent with the proposed inhibitoryeffect of glucocorticoids on collagen expression (as reviewed in Cooper,et al., 1999 J. Endocrinol. 163:159-164). As with other genes, ON and OPexpression did not appear to be affected by induction condition.Moreover, osteogenic induction resulted in significant decreases in OPlevel, as measured by microarrays, and was consistent with decreasesobserved upon induction of rat bone marrow stromal cells (Malaval, etal., 1994 J. Cell. Physiol. 158:555-572). While not restricted toosteogenic cells, both OP and ON are found in high amounts in bonetissue. Therefore, their expression, together with osteoblast-specificgenes like OC, supports the osteogenic capacity of PLA cells. Inaddition to these osteogenic genes, osteo-induced PLA cells expressedseveral other genes, including the proteoglycans decorin and biglycanand the transcription factors PPARγ1 and PPARδ.

[0347] In addition to the RT-PCR results, expression of several proteinscharacteristic of osteogenic differentiation was also observed usingboth IF and Western blotting. In support of the RT-PCR data, control andosteo-induced PLA cells expressed several proteins consistent with anosteogenic phenotype, including CNI, decorin, biglycan, OP and ON.Significant differences in CNI, decorin, biglycan and ON expression werenot observed upon osteogenic induction and an increase in OP expressionwas seen after 3 weeks induction. Expression of ON and OP was alsoobserved in control and osteo-induced PLA cells using IF withdifferences in intracellular expression pattern detected between the twocell populations. Specifically, OP expression in both control andinduced PLA cells concentrated to a perinuclear location, while itsdistribution appeared to be more uniform in MSC samples. Thisperinuclear concentration has been observed in MSCs during osteogenesisand is a characteristic of secreted proteins (Zohar, et al., 1998 Eur.J. Oral Sci. 106:401-407). However, contrary to this study, a definedperinuclear concentration of OP was not observed in our MSC populationsand may represent a clonal variant or specific culture conditions.Rather, OP in the MSCs concentrated to the cell surface and at cellprocesses. This focal distribution has also been observed in MSCs andmay indicate cell migration by these cells during differentiation(Zohar, et al., 1998 Eur. J. Oral Sci. 106:401-407). Similarintracellular patterns were observed for ON in control PLA and MSCsamples. In these cells, ON was distributed throughout the cell in afine punctate pattern and a low level was also found in the nucleus.Osteogenic induction did not alter this pattern in MSCs. However, thenuclear expression was lost upon differentiation of PLA cells.Furthermore, while ON was found in virtually all control PLA cells, notall osteo-induced PLA cells were ON-positive. Rather, expression of thisprotein was found in regions of high cell density. Finally, noexpression of OC was observed in control PLA cells, whereas a very lowlevel was detected in undifferentiated MSCs, consistent with the RT-PCRfindings. Osteogenic induction resulted in OC expression by a smallpercentage of the osteogenic PLA cells, while a larger percentage ofosteogenic MSCs expressed this protein. The expression pattern of OC wassimilar in both osteogenic PLA and MSCs: distributed throughout the celland concentrated at defined regions along the cell surface. Together,with CNI, OP and ON, the expression of OC is supportive of the RT-PCRdata and further confirms the osteogenic capacity of PLA cells in vitro.

PLA Cells Undergo Adipogenic Differentiation

[0348] The differentiation of adipocytes in culture is dependent uponmany factors, including serum, hormonal supplementation (insulin) andpharmacologic agents (indomethacin, IBMX) (Green, et al., 1974 Cell3:127-133; Russell, TR 1976 Proc. Natl. Acad. Sci. USA 73:4516-4520;Williams and Polakis 1977 Biochem. Biophys. Res. Commun. 77:175-186).However, initiation of the adipogenic program, in contrast to terminaldifferentiation, does not require such adipogenic agents but may bedependent upon increased culture confluence. Moreover, it is known thatreversible growth arrest at confluence must occur before mostpre-adipocytes can commit to the adipogenic lineage (Scott, et al.,1982, J. Cell Biol. 94:400-405; Speigelman and Farmer 1982 Cell29:53-60; Trayhurn and Ashwell 1987 Proc. Nutr. Soc. 46:135-142). Asadipogenic differentiation proceeds, a loss of proliferative potentialis observed and the irreversible loss of replication potential is acharacteristic of terminal adipocyte differentiation. To investigate ifPLA cells exhibit the same characteristics, PLA proliferation wascorrelated to adipogenesis, as measured by Oil Red O accumulation.Consistent with studies on pre-adipocyte cell lines, high levels ofdifferentiation occurred in confluent PLA cultures. Differentiating PLAcells assumed a more expanded morphology and began to accumulateintracellular lipid droplets as early as 2 weeks induction .Differentiation proceeded with no significant increase in PLA cellnumber, suggesting that cell number and growth kinetics are linked toPLA adipogenesis (FIG. 29).

PLA—Cd Markers and ECM—Supplements

[0349] Adipogenic differentiation is accompanied by several molecularand biochemical events, including the increase in lipogenic enzymes thatcatalyze the conversion of glucose into fatty acids and triglycerides.Glycerol-3-phosphate (G3P) is the primary substrate for triglyceridesynthesis in adipose tissue and the adipose conversion of 3T3 cells ischaracterized by a dramatic increase in the enzymatic source of G3P,glycerophosphate dehydrogenase (GPDH) (Kuri-Harcuch, et al., 1978 J.Biol Chem. 252:2158-2160; Pairault Greem 1979 J. Biol. Chem. 76). Basedon this, GPDH activity was measured in adipo-induced PLA and 3T3-L1cells. No significant difference in GPDH levels was detected betweendifferentiated cells and non-induced controls until 3 weeksdifferentiation. Moreover, the initial period of differentiation wasassociated with higher basal GPDH levels. The increased level of GPDH inadipo-induced PLA cells was associated with the appearance of Oil Red Ostaining (FIG. 29). Induction from 3 to 4 weeks resulted in asignificant increase in GPDH in both differentiated PLA cells and 3T3-L1controls and coincided with increased lipid accumulation. Continueddifferentiation for an additional week did not significantly changeenzyme levels in these cell populations. A similar pattern of GPDHactivity was also observed in adipo-induced MSCs. Therefore, theincrease in GPDH enzyme activity in PLA cells induced toward theadipogenic lineage indicates that these cells may be undergoingadipogenic differentiation.

[0350] Like osteogenesis, adipogenesis is characterized by theexpression of a distinct set of genes that are involved in lipidsynthesis and storage. One of these genes, PPARγ2, is a member of thePPAR nuclear hormone receptor superfamily, together with PPARγ1 andPPARδ (reviewed in (Fajas, et al., 1998 Curr. Biol. 10:165-173). PPARγ2has been identified as part of a heterodimeric complex (with ARF6 andthe retinoid X receptor) that acts as a key transcriptional regulator ofthe tissue-specific aP2 gene (Totonoz, et al., 1995 Nucl. Acid Res.).Moreover, PPARγ2 is expressed at high levels specifically in fat and isinduced early in the differentiation of cultured adipocyte cell lines(Totonoz, et al., 1994 Genes Dev. 8:1224-1234; Totonoz, et al., 1994Cell 79:1147-1156). Consistent with this, PPARγ2 was specificallydetected in adipo-induced PLA and MSC samples. Initial differentiation(i.e. 4 days) of these cell populations was characterized by the absenceof this transcription factor and is agreement with previous results fromdifferentiating 3T3 adipocytes (Totonoz, et al., 1994 Genes Dev.8:1224-1234; Totonoz, et al., 1994 Cell 79:1147-1156). Detectable levelsof PPARγ2 were observed after one week induction. However, expressionlevels were significantly higher at this time point in adipo-induced PLAcells, suggesting that the kinetics of PPARγ2 expression may differslightly between MSC and PLA populations. Distinctions in PPARγ1expression were also observed between PLA cells and MSCs. A similartime-dependent increase in PPARγ1 expression was observed in PLA cellsand MSCs. However, early differentiation (i.e. 4 days) of MSCs wasassociated with an absence of this transcription factor while low levelswere observed in induced PLA cells. Moreover, no PPARγ1 was detected incontrol MSCs. Finally, while detectable levels of PPARγ1 were seen innon-induced PLA cells, adipogenic induction was associated with asignificant increase in expression, consistent with adipogenicdifferentiation. PPARγ2 is associated with growth arrest and earlycommitment of pre-adipose cells to the adipogenic lineage. This periodof differentiation also marks the point at which the gene LPL isexpressed (Ailhaud, et al., 1992 Annu. Rev. Nutr. 12:207-233; Fajas, etal., 1998 Curr. Biol. 10:165-173). LPL (Lipoprotein Lipase) isubiquitously expressed but is significantly upregulated in adiposetissue. Through its hydrolysis of triglycerides, LPL promotes theexchange of lipids and affects the metabolism of severaltriglyceride-rich lipoproteins, including HDL and LDL (Eisenberg, etal., 1984 J. Lipid Res. 25:1017-1058). Consistent with its ubiquitousexpression, non-induced PLA and MSC controls expressed a low level ofLPL. However, adipogenic induction of both PLA cells and MSCs wasassociated with a significant increase in the expression of this gene.This increase was observed after one week induction and levels remainedequivalent throughout the remaining differentiation period. Finally,extended differentiation of preadipocytes results in the expression ofthe late adipogenic markers and is associated with the accumulation oflipid within the maturing adipocyte (Ailhaud, et al., 1992 Annu. Rev.Nutr. 12:207-233; Fajas, et al., 1998 Curr. Biol. 10:165-173). One suchlate marker is the fatty acid binding protein, aP2 (Bemlohr, et al.,1984 Proc. Natl. Acad. Sci. USA 81:468-472; Bernlohr, et al., 1985Biochem. Biophys. Res. Comun. 132:850-855). Consistent with previousresults (Bemlohr, et al., 1985 Biochem. Biophys. Res. Comun.132:850-855), aP2 was detected in 3T3-L1 controls, along with LPL andPPARγ2. However, despite its classification as a late marker inadipocytes, aP2 expression was observed throughout adipogenic inductionin both PLA cells and MSCs and levels appeared to be equivalent at eachinduction point. Moreover, aP2 expression preceded that of PPARγ2, indirect contrast to the pattern of expression observed in pre-adipocytedifferentiation ((Totonoz, et al., 1994 Genes Dev. 8:1224-1234; Totonoz,et al., 1994 Cell 79:1147-1156). Consistent with its function inadipogenesis, extremely low levels of aP2 were found in non-inducedcontrols. This constitutive expression was in agreement with theexpression of aP2 in tissues other than fat (Zezulak and Green 1985 1985Mol. Cell Biol. 5:419-421) and is similar to the LPL results. Takentogether, the adipogenic-specific expression of PPARγ2 in adipo-inducedPLA cells, together with the upregulated expression of LPL and aP2 issupportive of the adipogenic capacity of these cells.

[0351] Furthermore, the adipogenic capacity in combination with theosteogenic potential of these cells suggests that PLA cells may possessmulti-lineage potential.

PLA Cells Undergo Chondrogenesis

[0352] Chondrogenic differentiation of cell lines requires high densityculture (Johnstone, et al., 1998 Exp. Cell Res. 238:265-272),duplicating the process of cellular condensation (Fell1925 J. Morphol.Physiol. 40), in addition, to supplementation with specific growthfactors, such as TGFβ1, TGFβ3 or BMP2 (Johnstone, et al., 1998 Exp. CellRes. 238:265-272; Mackay, et al., 1998 Tissue Eng. 4:415-428).Consistent with this, aggregate culture of PLA cells in CM, containingTGF-62 1, resulted in the formation of small, compact micromass nodulesas early as 24 hours induction . Induced PLA nodules stained positivelyusing the stain Alcian Blue, consistent with the presence of sulfatedproteoglycans within the nodule ECM and in agreement with the resultsdescribed in Example 7 above. Alcian blue staining appeared toconcentrate more in the interior of the nodule and was apparent as earlyas 3 days induction. Consistent with Alcian Blue staining, PLA nodulesalso contained keratan- and chondroitin-4-sulfate, two proteoglycansexpressed in high amounts in cartilage. In support of these results, theexpression of KS and CS has also been observed in human bone marrow MSCsinduced toward the chondrogenic lineage (Yoo, et al., 1998 J. Bone JointSurg. Am 80:1745-1757: Yoo, et al., 1998 Clin. Orthop. S73-81). Inaddition to sulfated proteoglycans, PLA nodules also expressed collagentype II, a collagen isoform characteristic of cartilage tissue. Finally,PLA nodules cultured under high-density conditions and maintained innon-inductive control medium did not form nodules and failed to stainfor any cartilage-specific histologic marker, thus confirming thespecificity of our induction conditions.

[0353] Quantitation of sulfated proteoglycans can be accomplished usinga metachromatic dimethyldimethylene blue assay (Famdale, et al., 1986Biochimica et Biophysica Acta 883:173-177). Consistent with ourimmunohistochemical results, the DMMB assay confirmed the presence ofsulfated proteoglycans in the differentiated PLA samples.

[0354] Moreover, a time-dependent increase in KS and CS withinchondrogenic PLA nodules was observed up to 2 weeks of induction. Asimilar increase has also been observed in induced MSC cultures (Yoo, etal., 1998 J. Bone Joint Surg. Am 80:1745-1757: Yoo, et al., 1998 Clin.Orthop. S73-81) and suggests that PLA cells have accumulated an ECMcharacteristic of cartilage tissues. PG levels decreased slightly beyond2 weeks induction and may represent remodeling of the cartilagenous ECM.Non-induced PLA cells, maintained under high-density conditions, werealso associated with an ECM containing these proteoglycans. Moreover,basal PG levels were greater than induced PLA sample at 4 and 7 days.However, significantly more proteoglycan accumulation was observed ininduced PLA nodules at days 14 and 21. The significant accumulation ofKS and CS within the ECM of induced PLA nodules, together with thehistological results suggests that PLA cells also possess in vitrochondrogenic capacity when cultured under high-density conditions.

[0355] Induction of PLA cells in CM resulted in the expression ofseveral genes consistent with chondrogenesis. CNII expression wasobserved specifically in induced PLA cells and was restricted to day 7and 10 and supported our immunohistochemical results. A restrictedexpression pattern similar to CNII was observed in PLA nodules usingprimers designed to the amino terminus of aggrecan (AG), a largeproteoglycan expressed in high amounts in cartilage. Expression ofaggrecan was also observed in PLA samples using primers to the carboxyterminus (PG). However, in addition to expression at days 7 and 10, PGwas also detected at day 14 in these nodules. In support of the PLAresults, expression of aggrecan in induced NHCK nodules was detectedusing both amino and carboxy primer sets. Like CNII, the expression ofaggrecan was specific to induced PLA and NHCK nodules. In addition toCNII, chondrogenic induction of PLA cells resulted in the restrictedexpression of CNX, a marker of hypertrophic chondrocytes. Expression ofCNX was detected at day 14 and suggests that PLA nodules undergohypertrophy over time. Induced NHCK samples also expressed CNX, althoughat a lower level. PLA nodules were also associated with additionalcollagen types, including CNI and CNIII. While the majority of PLAsamples examined exhibited a restricted collagen pattern (day 4 only),CNI and was detected in a few PLA samples up to day 14. The expressionof CNI has also been observed in human MSC nodules by fibroblastic cellslocated in the outer nodule, leading researchers to suggest that thisregion is comprised of perichondrium-like cells involved in thedifferentiation process (Yoo, et al., 1998 J. Bone Joint Surg. Am80:1745-1757: Yoo, et al., 1998 Clin. Orthop. S73-81). In support ofthis, perichondrium-like cells have also been observed in high-densityembryonic chick limb-bud cell cultures and cell aggregates (Osdoby andCaplan 1979 Devel. Biol. 73:84-102; Tachetti, et al., 1987 J. Cell Biol.106:999-1006). Therefore, the continued expression of CNI in select PLAsamples may be due to the presence of a similar cell population.

[0356] Induced and control PLA cells also expressed the proteoglycans,decorin and biglycan and the gene CBFA1. Decorin and biglycan make upthe majority of the small leucine-rich proteoglycans within thecartilagenous ECM and their expression within PLA nodules furthersupports the chondrogenic phenotype. In addition to its expressionduring osteogenesis, a role for CBFA-1 in the hypertrophy and terminaldifferentiation of chondrocytes has recently been confirmed (Enomoto, etal., 2000 J. Biol. Chem. 275:8695-8702. Therefore, the expression ofCBFA-1, together with CNX, may indicate terminal differentiation of PLAcells within the nodule. Chondrocyte hypertrophy may also precede theossification of cartilagenous tissue. However, expression ofbone-specific OC by chondrogenic PLA or NHCK cells was not seen at anytime point, confirming the absence of osteogenic differentiation withinthe PLA nodule. Interestingly, micromass culture of MSCs in CM did notresult in the formation of nodules and was not examined. Taken together,the specific expression of CNII, aggrecan and CNX in induced PLAnodules, in addition to the presence of keratan- andchondroitin-4-sulfate within the ECM supports the chondrogenic phenotypeof these cells. Moreover, the chondrogenic capacity of PLA cells,together with their osteogenic and adipogenic potential, furthersupports the multi-lineage capacity of these putative stem cells.

PLA Cells Undergo Myogenic Differentiation

[0357] RT-PCR analysis of PLA cells induced toward the myogenic lineageconfirmed the expression of several myogenic genes, including thetranscription factors MyoD1, myogenin and myf5, in addition to themuscle-specific protein, the myosin heavy chain. Determination of themyogenic lineage is thought to be controlled at the transcriptionallevel by MyoD1 and myf-5, which are expressed in proliferating myoblasts(Atchley, et al., Proc. Natl. Acad. Sci. 91:11522-11526; Lassar, et al.,1994 Curr. Opin. Cell Biol. 6:432-442; Weintraub, et al., 1994 GenesDev. 15:2203-2211), whereas execution of the myogenic differentiationprogram is controlled by myogenin and MRF4 expression (emerson, et al.,1993 Curr. Opin. Genet. Dev. 3:265-274; Olson, et al., 1996 Cell 5:1-4).Finally, terminal differentiation of myoblasts can be confirmed throughthe expression of the myosin heavy chain. Consistent with thesefindings, the expression of myf5 was restricted to the first 3 weeks ofmyogenic PLA induction while increased MyoD1 expression was detectedwithin the first week relative to the remainder of the differentiationperiod. Myo-induced PLA cells also expressed myogenin at relativelyequivalent levels throughout the 6 week induction period. Finally,increased expression of the myosin heavy chain was detected at 6 weeksinduction and suggests that PLA cells underwent of terminaldifferentiation. The expression of myf5 and myogenesis further supportsthis potential and, together with the osteogenic, adipogenic andchondrogenic capacity of PLA cells, indicates their potential fordifferentiation to multiple mesodermal lineages.

PLA Cells May Possess Neurogenic Potential

[0358] True pluripotency of a stem cell is achieved upon differentiationto cells from distinct embryologic lineages. Recent reports havedocumented the differentiation of MSCs to neural cells (Deng, et al.,1994 Genes Devel. 8:3045-3057; Kopen, et al., 1999 Proc. Natl. Acad.Sci. USA 95:3908-3913; Sanchez-Ramos, et al. Exp. Neurol. 164:247-256;Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370) and neural stemcells (NSCs) to haematopoietic cells (Bjornson, et al., 1999 Science283:534-537), suggesting that stem cell populations may not be asrestricted as previously thought. Based on these findings, weinvestigated if PLA cells could be induced beyond their putativemultilineage mesodermal capacity. To this end, PLA cells were culturedin a medium known to induce neurogenic differentiation (Vescovi, et al.,1999 Exp. Neurol. 156:71-83; Woodbury, et al., 2000 J. Neurosci. Res.61:364-370) and differentiation assessed by staining for neural markers,including NSE, trk-a and MAP-2 or for the expression of GFAP and GaIC,markers of astrocytes and oligodendrocytes, respectively. Themorphologic and histologic data suggest that PLA cells, like MSCs,possess neurogenic potential in vitro. Induction of PLA cells in NM fora minimum of 30 minutes resulted in a dramatic change in morphology withcells assuming a neuronal-like phenotype. NM-induced PLA cells underwentretraction, forming compact cells bodies with multiple extensions. Cellbodies became more spherical and cell processes exhibited secondarybranches with increasing induction time. A time-dependent increase inthe proportion of PLA cells with this phenotype was observed in allinduced PLA cultures. Similar morphologic changes have been observedupon neurogenic induction of MSCs from both rodents and human (Woodbury,et al., 2000 J. Neurosci. Res. 61:364-370). Moreover, this PLAmorphology was similar to that observed upon NGF stimulation of PC 12cells, a neuroendocrine cell line similar to primary sympathetic neurons.

[0359] The observed morphologic changes in neuro-induced PLA cells wereaccompanied by the increased expression of neuron-specific markers, suchas NSE, trk-a and NeuN, and did not result in expression of markers forastrocytes and oligodendrocytes. Furthermore, expression of thesemarkers was also observed in PC12 cultures, suggesting that PLA cellsmay be assuming a neuronal-like phenotype. In support of the PLAresults, increased expression of NSE, a neuron-specific enolase, andtrk-a has been observed upon induction of MSCs with β-ME, withapproximately 100% of the neuronal-like MSCs positive for these markers(Woodbury, et al., 2000 J. Neurosci. Res. 61:364-370). Like the MSCstudies, all PLA cells exhibiting a neuronal phenotype expressedsignificant levels of NSE and trk-a. In addition to NSE, expression ofNeuN has also been used to identify neuronal development in neurogenicprecursors and MSCs (Sanchez-Ramos, et al., 2000 Exp. Neurol.164:247-256). Specifically, NeuN is expressed in post-mitotic neurons(Sarnat, et al., 1998 Brain Res. 20:88-94) and its appearance is thoughtto coincide with the withdrawl of the developing neuron from the cellcycle and/or the initiation of terminal differentiation (Mullen, et al.,1992 Development 116:210-211). The expression of NeuN within theneuronal-like PLA cells, together with the presence of NSE and trk-a,further supports the development of a neuronal phenotype in PLA cells.Moreover, the expression of NeuN may indicate the development of apost-mitotic neuronal phenotype. In contrast to NSE, trk-a and NeuN,expression of the mature neuronal markers, tau, MAP-2 and NF-70, was notobserved , suggesting that induced PLA cells represent an earlydevelopmental stage. Consistent with this, MAP-2 expression in inducedMSC cultures has not been observed by several groups and may reflect theinduction conditions used or the need for prolonged induction time(Deng, et al., 2001 Biochem. Biophys. Res. Commun. 282:148-152;Sanchez-Ramos, et al., 2000 Exp. Neurol. 164:247-256).

[0360] Finally, the putative neuronal potential of PLA cells wasconfirmed using RT-PCR. Consistent with the immunohistochemistryresults, no expression of GFAP could be detected, supporting therestriction of induced PLA cells to the neuronal lineage. In addition,PLA cells were examined for the expression of the gene nestin. Nestin,an intermediate filament protein, has been detected in high amounts inCNS stem cells (Lendahl, et al., 1990 Cell 60:585-595), within thedeveloping neural tubes of mice (Frederikson and McKay 1988 J. Neurosci.8:1144-1151) and in MSCs induced toward the neurogenic lineage(Sanchez-Ramos, et al., 2000 Exp. Neurol. 164:247-256). Differentiationof neural precursors results in a decrease in nestin expression levels,indicating that this protein can be used as a marker of a progenitorphenotype (Johe, et al., 1996 Genes Dev. 10:3129-3140; Lendahl, et al.,1990 Cell 60:585-595). The expression of nestin in control PLA culturesis supportive of the presence of neurogenic precursors within the PLA.However, differentiation of PLA cells did not result in an appreciabledecrease in nestin expression. This may be due to two possibilities: 1)the differentiation of PLA cells into a neurogenic progenitor populationonly or 2) the differentiation of PLA cells into an early neuronal-likecell that retains nestin expression. In support of the latter, nestinexpression was also detected in NGF-treated PC12 controls. Based onthis, together with the expression of NeuN, NSE and trk-a in induced PLAcells, leads us to favor the latter possibility and further studies arewarranted. Like our IH results, RT-PCR analysis failed to detectexpression of a mature neuronal marker (GAD65), a marker detected inPC12 controls and brain. It is possible that additional growth factorsor a prolonged induction period may be required to induce PLA cells intoa more mature stage. Finally, induction of PLA cells with NM appeared torestrict their development to cells characteristic of the CNS, as thecells did not express ChaT, a specific marker of peripheral nerves.Nestin expression has also been observed in non-induced MSCs, inaddition to myogenic cells, newly formed endothelial cells, epithelialcells of the developing lens and hepatic stellate cells. This broaddistribution indicates that nestin cannot be used as a neurogenicprecursor marker per se. However, combined with the expression ofadditional neuronal markers, such as NeuN, the possibility that PLAcells are forming precursors of the neuroectodermal lineage isstrengthened.

ADSC Clonal Isolates Demonstrate Multi-Lineage Capacity

[0361] Multi-lineage differentiation by PLA cells may result from thecommitment of multiple lineage-specific precursors rather than thepresence of a pluripotent stem cell population within adipose tissue.Therefore, multi-lineage differentiation by clonal isolates derived fromsingle PLA cells is critical to the classification of PLA cells as asource of stem cells. In support of this, single PLA cell isolatesexpanded in culture exhibited multi-lineage capacity in vitro, stainingpositively for alkaline phosphatase (osteogenesis), Oil Red O(adipogenesis) and Alcian Blue (chondrogenesis). Clonal analysisresulted in the isolation of several lineage combinations, includingtri-lineage (osteogenic, adipogenic and chondrogenic), dual-lineage(osteogenic/adipogenic, osteogenic/chondrogenic) and single lineage(adipogenic only). The tri-lineage clones were subsequently termedAdipose Derived Stem cells (ADSCs) and were analyzed for multilineagepotential using RT-PCR. Consistent with multilineage capacity ADSCsexpressed several genes characteristic of osteogenesis (OC, ON, OP, CNIand AP), adipogenesis (PPARγ2, aP2 and LPL) and chondrogenesis (AGG,CNX, decorin and biglycan). Furthermore, several tri-lineage ADSCs alsoexpressed the neuronal marker trk-a using IH (FIG. 39). Based on theseresults, the expression of multiple lineage-specific mesodermal genes byADSCs suggests that these isolated clones possess multipotentiality andmay be considered stem cells.

EXAMPLE 12

[0362] The following provides a description of molecular and biochemicalcharacterization of adipose-derived stem cells.

Materials and Methods

[0363] All materials were purchased from Sigma (St. Louis, Mo.), VWR(San Dimas, Calif.) and Fisher Scientific (Pittsburgh, Pa.) unlessotherwise stated. All tissue culture reagents were purchased from LifeTechnologies (New York, N.Y.). Fetal Bovine Serum (FBS) and Horse Serum(HS) were purchased from Hyclone (Logan, Utah) and Life Technologies,respectively.

Antibodies

[0364] Monoclonal antibodies to CD29, CD34, CD44, CD45, CD58, CD90,CD104, CD105 and CD140a were purchased from Pharmingen (Bedford, Mass.).Monoclonal antibodies to CD31 and CD71 were obtained from R&D Systems(Minneapolis, Mass.) and Zymed (S. San Francisco, Calif.), respectively.FITC and PE-conjugated anti-CD antibodies used for flow cytometry (FC)were purchased from Pharmingen. A monoclonal antibody to the SH3 antigenwas produced from the SH3 hybridoma (ATCC). The Stro-1 hybridomasupernatant was the generous gift of Dr. John Fraser (UCLA). Monoclonalantibodies to the human collagens 1 (αCNI) and 4 (αCNIV) were purchasedfrom Sigma. Monoclonal antibodies to human collagen 3 (αCNIII) andcollagen 5 (αCNV) was purchased from Biogenesis (Kingston, N.H.).

Cell Harvest Culture and Differentiation Conditions

[0365] Processed lipoaspirate (PLA) cells were obtained from rawlipoaspirates and cultured as described previously (Zuk, P. et al., 2001Tissue Engineering 7:209-226). PLA cells were maintained innon-inductive Control medium (Table 13) while MSCs were maintained inspecialized Control medium (Clonetics). PLA cells were induced towardthe desired mesenchymal lineages using the induction media outlined inTable 13. MSCs were induced using the commerical control mediumsupplemented with the same growth factors as outlined in Table 13.

PLA Clonal Isolation and Analysis: Adipose-Derived Stem Cells (ADSCs)

[0366] ADSC Isolation: PLA cells were plated at extremely low confluencein order to result in isolated single cells. Cultures were maintained inControl medium until proliferation of single PLA cells resulted in theformation of well-defined colonies. The single PLA-cell derived colonieswere termed Adipose Derived Stem Cells (ADSCs). ADSCs were harvestedusing sterile cloning rings and 0.25% trypsin/EDTA. The harvested ADSCswere amplified in Cloning Medium (15% FBS, 1% antibiotic/antimycotic inF12/DMEM (1:1)).

Indirect Immunofluorescence

[0367] Indirect Immunofluorescence (IF): PLA cells, ADSCs and MSCs wereprocessed for IF as described previously (Zuk, P. et al., 2001, TissueEngineering, 7:209-226) using the anti-CD marker antibodies outlined inTable 14. In addition, PLA cells were incubated with supernatantsproduced from the STRO-1 and SH3 hybridoma cell lines. To determine thecell characteristics of differentiated PLA cells and MSCs, cells wereinduced toward either the osteogenic lineage for 3 weeks or theadipogenic lineage for 2 weeks and incubated with anti-CD antibodies.The differentiated cells were also analyzed using antibodies to humancollagens 1, 4 and 5.

Flow Cytometry

[0368] PLA cells from multiple donors, in addition to MSCs, werecultured for 3 weeks in Control medium and analyzed for the expressionof CD antigens by flow cytometry (FC) as described previously (Zuk, P.et al., 2001, Tissue Engineering, 7:209-226). PLA cells were alsoinduced in either OM or AM for 2 weeks prior to analysis. Briefly, cellswere harvested a 80% confluence with trypsin/EDTA, washed andresuspended in Flow Cytometry Buffer (FCB) at a concentration of 1×10⁶cells/ml. One hundred microliters of the cell preparation (1×10⁵ cells)were stained with saturating concentrations of FITC-conjugated (anti-CD14, 44, 45 61, 71, 90 and 105) or PE-conjugated (anti-CD 13, 16, 31, 34,44, 49d, 56, 62E and 106) antibodies for 1 hour at 4° C. Cells were alsoincubated with isotype-matched IgG's as a control to assessautofluorescence. After incubation, the cells were washed three timeswith FCB and resuspended for analysis. Flow cytometry was performed on aFACStar flow cytometer (Becton Dickson). The geometric means, calculatedfrom the absolute numbers of cells per 10,000 events are shown in Table12.

Results PLA Cells Share Many Similarities with MSCs

[0369] The results described in this example demonstrate themutli-lineage potential of adipose-derived stem cells and their clonalisolates. In order to characterize the PLA population further, cellswere examined using indirect IF and FC and compared to a commercialpopulation of human MSCs. MSCs have been shown to express a unique setof cell surface markers that can be used to help identify this stem cellpopulation (Table 14) (Bruder, S. P. et al., 1998, Clin. Orthop.,S247-256; Conget, P. A. et al., 1999, J. Cell Physiol, 181:67-73;Pittenger, M. F et al., 1999, Science, 284:143-147.) Like MSCs, PLAcells expressed several of these proteins (FIGS. 23 and 24), supportingthe characterization of these cells as stem cells. Approximately 100% ofthe PLA and MSC cultures were positive for the expression of CD29, CD44,CD90 and CD105/SH2 with high expression levels for each of these markersbeing observed in both cell populations. Both cell populations alsoexpressed the SH3 antigen, which, together with SH2, is considered aspecific marker for MSCs (Haynesworth, S. E. et al., 1992, Bone,13:69-80.)

[0370] In addition, the majority of PLA cells and MSCs were alsopositive for the transferrin receptor, CD71, indicating that a fractionof these cell populations were replicating. PLA and MSCs did not expressthe haematopoietic lineage markers, CD31 and CD34. A small number of PLAsamples did show negligible staining for CD45, although the number ofCD45-positive cells did not exceed 5% of the total PLA cell number.Unlike MSCs, no staining for the adhesion molecule CD58 was observed inPLA cells. The IF results were subsequently confirmed by FC (FIG. 24,Panel B). Both MSC and PLA cells showed similar profiles, comprisedmainly of a population of relatively small, agranular cells (FIG. 24,Panel A). However, a greater proportion of the PLA population did appearto contain larger, granular cells (see upper right corner), while alarger proportion of the MSC population contained smaller agranularcells. FC confirmed the expression of CD44, CD71, CD90 and CD105 on bothPLA and MSCs and did not detect significant levels of CD3 1, CD34, CD45and CD104. In addition to these markers, FC also measured expression ofCD13, CD49d, SH3 and STRO-1 on PLA cells yet did not detect expressionof CDs 4, 8, 11, 14, 16, 19, 33, 56, 62E and 106 (Table 12). Takentogether, the immunofluorescent and flow results demonstrate severalsimilarities in CD expression profiles between PLA populations and bonemarrow-derived MSCs.

Phenotypic Characterization of Differentiated PLA Cells

[0371] Differentiation of stem cells may alter the expression of severalcell surface and intracellular proteins. In order to characterizedifferentiated PLA cells, cells from the same patient were maintained innon-inductive Control medium or were induced toward the osteogenic andadipogenic lineages. Control and differentiated PLA cells weresubsequently analyzed by IF and compared to MSCs as a control. Theresults are presented in Table 15. PLA cells induced for 3 weeks in OMunderwent increased proliferation and did not show any significantdifferences in CD marker profile when compared to undifferentiated PLAcells. Like control PLA cells, expression of CD45 was not observed inosteogenic PLA cells while significant expression of CD44 and CD90 wasdetected (FIG. 40, Panel A: PLA—Bone).However, in contrast to controlcells, osteogenic differentiation resulted in localized areas of CD34expression. Like PLA cells, the CD marker profile of control andosteo-induced MSCs was similar, with the exception of CD34 and CD45. Asshown in FIG. 40, Panel B, expression of CD34 and CD45 was not observedin control MSCs. However, a slight increase in CD34 expression level wasobserved upon induction in OM while an increased number of CD45-positivecells were detected.

[0372] To induce adipogenic differentiation, PLA cells were maintainedfor a minimum of 2 weeks in AM. In order to correlate CD markerexpression to cell morphology, fluorescent micrographs were overlaidwith light micrographs (inset pictures). Induction of PLA cells with AMresulted in an expanded cellular morphology and the accumulation ofmultiple, intracellular lipid vacuoles, consistent with adipogenesis.These lipid-containing PLA cells were considered to be mature adipocytes(white arrows) (FIG. 41, Panel A: PLA—Fat). Like osteogenesis,adipogenic differentiation appeared to result in slightly increased CD34levels in both fibroblastic and lipid-containing PLA cells (FIG. 41,Panel A). In addition, a negligible fraction of the adipogenic PLAcultures contained CD45-positive cells. However, these cells did notcontain the lipid vacuoles characteristic of mature adipocytes(CD45—inset). A significant level of CD44 was also detected inadipogenic PLA cultures. However, lipid-filled PLA cells appeared toexpress lower levels of CD44 in comparison to their fibroblasticcounterparts (open arrows—CD44−ve adipocytes, filled arrows—CD44+vecell). Furthermore, CD44 staining levels varied among the fibroblasts,ranging from intense to little or no CD44 expression. A similarrestriction was also observed for CD90 with all fibroblasts expressingthis protein at comparable levels. Like PLA cells, adipo-induced MSCsexpressed CD44 and CD90 and showed increased staining for CD34 and CD45(FIG. 41, Panel B and Table 16). However, unlike adipo-induced PLAcells, both fibroblastic and lipid-filled MSCs (filled vs. open arrows,respectively) appeared to express CD44 and CD90 at similar levels.

[0373] In order to confirm the immunofluorescent results, FC wasperformed on non-induced and differentiated PLA cells and the geometricmeans calculated for each CD marker protein (FIG. 42) (Table 16).Osteogenic differentiation did not appreciably change the size andgranularity of the PLA populations (FIG. 42, Panel A). Adipogenesis,however, resulted in a significant increase in the size and granularityof the PLA population, likely a reflection of the expanded cellularmorphology and the formation of intracellular lipid vacuoles. Consistentwith the immunofluorescent results, control PLA cells were negative forCD34 and CD45 expression (Panel B), nor could expression of CD14, CD16,CD31, CD34, CD45, CD56, CD61, CD62, CD105 or CD106 be measured in thesecells (Panel B). Differentiation appeared to increase CD34 and CD61expression with a greater increase being observed upon osteogenicdifferentiation. The increased CD34 expression was consistent with theincreased staining observed upon IF processing (FIG. 40). Slightincreases in CD56 and CD49d were detected specifically in osteo-inducedPLA cells. The expression of CD56 in adipo-induced PLA cells did notdiffer significantly from controls, while a decrease in CD49d expressionwas detected upon adipogenesis. FC also confirmed the expression ofCD13, CD44 and CD90 in control cells (FIG. 42, Panel C.). Osteogenesissignificantly increased expression of these markers and a furtherincrease in CD90 was measured in adipo-induced PLA cells. Finally,adipogenic differentiation resulted in a decreased expression of CD13and CD44 to below that of undifferentiated PLA cells. The decrease inCD44 was consistent with the IF results, in which lower expressionlevels were seen in lipid-containing PLA cells.

[0374] Mesodermally-derived cells, such as adipocytes and osteoblastsare associated with extensive extracellular matrices (ECMs). To assessthe expression of ECM proteins in differentiated PLA cells, adipogenicand osteogenic PLA cells were analyzed by IF for the expression of ECMcollagens. The results are summarized in Table 17. The majority ofundifferentiated PLA cells expressed collagen types 1 and 3 (CNI, CNIII)(FIG. 43, Panel A: PLA—Control). CNI and CNIII in fibroblastic PLA cellswere restricted to a defined perinuclear concentration and was evenlydistributed throughout while cells with an expanded morphology. Contraryto CNI and CNIII, the expression of collagen types 4 and 5 (CNIV, CNV)was restricted to defined culture regions of concentrated PLA cells andmatrix formation. The expression patterns of CNIV and CNV were fibrillarin nature, consistent with the secretion of these proteins into theextracellular space surrounding these cells.

[0375] Adipogenic differentiation of PLA cells inhibited both CNI andCNV expression (FIG. 43, Panel A: PLA—Fat). Differentiation alsoappeared to alter the intracellular expression pattern of CNIII,redistributing it evenly throughout the mature PLA adipocyte. Inaddition, a lower level of CNIV expression was observed in adipogenicPLA samples, with the majority of the CNIV fluorescence observed inlipid-filled PLA cells (see arrows; inset). While an inhibition of CNVexpression was also observed upon osteogenic induction, no significantdifference in CNI expression pattern could be detected for this lineage(FIG. 43, Panel A; PLA—Bone). Finally, osteogenesis appeared to increaseCNIV expression levels and resulted in a more widespread distribution ofthis collagen within the PLA culture. The expression patterns of CNI,CNIII, CNIV and CNV in control MSCs were found to be similar to controlPLA cells (FIG. 43, Panel B: MSC—Control). Like PLA cultures, CNIII andCNIV were detected in adipo-induced MSC samples. However, CNIV appearedto be restricted to extracellular fibrils rather than an intracellulardistribution (FIG. 43, Panel B: MSC—Fat, see arrows). In contrast toadipogenic PLA cultures, induction toward this lineage did not inhibitsynthesis of CNI and CNV by MSCs. Rather, these collagens could bedetected within extracellular fibrils and weak CNI expression could alsobe observed within lipid-filled MSCs. Finally, in contrast toosteo-induced PLA cells, osteogenic induction of MSCs did not alter theintracellular expression pattern of CNI or the synthesis andextracellular deposition of CNIV and CNV (FIG. 43, Panel B; MSC—Bone).Taken together, the immunofluorescent data suggests that both adipogenicand osteogenic differentiation of PLA cells leads to a remodeling of theassociated ECM, resulting in a matrix that appears to be distinct fromthose of MSCs.

PLA Clonal Isolates (ADSCs) Express a Similar Complement of CD MarkerProteins

[0376] Multi-lineage differentiation by PLA cells may be the result ofthe commitment of multiple lineage-specific precursors rather than thepresence of a pluripotent stem cell population within adipose tissue.Therefore, multi-lineage differentiation by clonal isolates derived fromsingle PLA cells is critical to the classification of PLA cells as asource of stem cells. In support of this, single PLA cells colonies,termed Adipose-Derived Stem Cells (ADSCs), exhibited multi-lineagecapacity in vitro (FIG. 35). Analysis of 500 ADSC isolates confirmeddifferentiation potential in approximately 6% of the total number ofclones examined. Seven ADSC isolates exhibited tri-lineage potential,differentiating into cells of the osteogenic, adipogenic andchondrogenic lineages, approximately 24% of the total number of ADSCspositive for differentiation potential (Table 11). Furthermore, aqualitative increase in differentiation level, as measured by histologicstaining, was also observed in these tri-lineage ADSC populations. Inaddition to tri-lineage ADSCs, several dual-lineage clones (O/A, O/C andA/O) and single adipogenic lineage clones were also isolated. Isolationand expansion of ADSCs did not alter the CD expression profile of theclones, as no difference in CD expression could be detected by IF.Furthermore, no difference was also observed between tri- and duallineage ADSC isolates (FIG. 36). Like the heterogenous PLA populations,ADSCs were positive for CD29, CD44, CD71 and CD90 expression, while noexpression of CD31, 34, 45 and 104 was observed. Therefore, the presenceof multi-lineage ADSC isolates within the heterogenous PLA cellpopulation and their identical CD marker profile to PLA cells furthersupports the theory that the adipose compartment is a source ofmulti-potential stem cells. TABLE 13 Lineage-specific differentiationinduced by media supplementation Medium Media Serum SupplementationControl DMEM 10% FBS None Adipogenic DMEM 10% FBS 0.5 mMisobutyl-methylxanthine (AM) (IBMX), 1 μM dexamethasone, 10 μM insulin,200 μM indomethacin, 1% antibiotic/antimycotic Osteogenic DMEM 10% FBS0.1 μM dexamethasone, 50 μM (OM) ascorbate-2-phosphate, 10 mM β-glycerophosphate, 1% antibiotic/ antimycotic

[0377] TABLE 14 Monoclonal antibodies to CD antigens: Reported cellspecificity and distribution CD Antigen Clone Cell Specificity 29Integrin β1 MAR4 broad distribution—lymphocytes, monocytes, granulocytesNOT on erythrocytes 31 PECAM-1 9G11 endothelial cells, platelets,monocytes, granulocytes, haematopoietic precursors 34 — 581 endothelialcells, some tissue fibroblasts, haematopoietic precursors 44 Pgp-1G44-26 leucocytes, erythrocytes, epithelial cells, platelets 45 LCA HI30leucocytes, haematopoietic cells 58 LFA-3 L306.4 widedistribution—haematopoietic cells, endothelial cells, fibroblasts 71 TfRH68.4 most dividing cells 90 Thy-1 5E10 immature CD34+ cells, cellscapable of long term culture, primitive progenitor cells 104  105 Endoglin — endothelial cells, B cell precursors, MSCs SH3 — —mesenchymal stem cells

[0378] TABLE 15 Immunofluorescent analysis of differentiated PLA and MSCpopulations CD marker PLA PLA-OM PLA-AM MSC MSC-OM MSC-AM CD29 +ve +ve+ve +ve +ve +ve CD31 −ve −ve −ve −ve −ve −ve CD34 −ve +/−, restricted+ve (weak) −ve +ve (weak) +ve (weak) CD44 +ve +ve +ve, +ve +ve +vefibroblastic cells CD45 −ve −ve +/−, −ve +ve +ve (weak) fibroblasticcells CD58 −ve −ve −ve +ve +ve +ve CD71 +ve +ve +ve +ve +ve +ve CD90 +ve+ve +ve +ve +ve +ve fibroblastic cells CD105 +ve +ve +ve +ve +ve +ve

[0379] TABLE 16 Flow cytometric analysis of CD marker expression inosteogenic, adipogenic and control PLA cells. CD Antigen PLA-CM PLA-OMPLA-AM CD13 148.88 924.79 134.34 CD14 2.43 3.54 3.08 CD16 2.38 3.43 2.70CD31 2.22 2.92 2.53 CD34 3.55 9.10 5.27 CD44 16.92 64.62 8.76 CD45 2.523.85 3.47  CD49d 5.33 13.05 4.27 CD56 2.66 4.86 2.72 CD61 3.68 7.55 4.12  CD62E 2.30 2.89 2.38 CD90 25.96 45.32 46.53 CD105  8.39 16.70 11.53CD106  2.45 3.27 2.51 SH3 8.95 25.15 14.65 −ve 2.59 3.57 3.08

[0380] TABLE 17 Immunuofluorescent staining patterns of extracellularmatrix collagens: Effect of differentiation. Collagen ImmunofluorescentStaining Pattern type PLA MSC PLA-Fat MSC-Fat PLA-Bone MSC-Bone 1punctate + punctate + no expression weak cellular expression +punctate + punctate + perinuclear perinuclear fibrillar patternperinuclear perinuclear concentration concentration concentrationconcentration 4 fibrillar, localized fibrillar pattern cellulardistribution, fibrillar, decreased fibrillar, weak fibrillar pattern todefined regions lipid-filled cells only expression expression 5fibrillar, localized fibrillar pattern no expression fibrillar patternno expression cellular + fibrillar to defined regions pattern

Discussion

[0381] In this study, a more comprehensive characterization of the PLAand ADSC populations was performed using a combination ofimmunofluorescence and flow cytometry. While PLA cells expressed asimilar complement of CD antigens with MSCs (positive: CD29, CD44, CD71,CD90, CD105 SH3, negative: CD31, CD34, CD45), the expression of CD58,CD104 and CD140a differed on PLA cells when examined byimmunofluorescence. Flow cytometry also confirmed the expression of CD13and the absence of CD14, CD16, CD56 and CD62E. Subtle distinctionsbetween non-induced and differentiation PLA cells could be determinedusing flow cytometry. Specifically, increases in CD13, CD44 and CD90were observed upon osteogenic induction, whereas CD13 and CD44 levels inadipogenic cultures were found to be lower. Consistent with this, IFanalysis indicated a lower level of CD44 expression within lipid-filledPLA cells (i.e. mature adipocytes). Osteogenic differentiation alsoresulted in slight increases in CD34, CD49d, CD56 and CD61. CD34expression was confirmed using immunofluorescence, with CD34-positiveregions being observed in osteogenic PLA cultures. ADSC clonalpopulations also expressed a similar complement of CD antigens to thatobserved in the heterogenous PLA population, suggesting that clonalisolation and expansion of these cells does not affect cell surfaceprotein expression. Finally, differentiation of PLA cells also resultedin changes to the associated ECM and differences in the expressionpatterns and levels of collagen types 1, 4 and 5 were found betweendifferentiated PLA and MSC cultures. Taken together, this data suggeststhat PLA cells may represent a stem cell population within adiposetissue but is a population that possesses subtle distinctions from MSCs.

[0382] While PLA cells expressed a similar complement of CD antigenswith MSCs, an established mesenchymal stem cell population, PLA cellsdid show subtle differences in the expression of CD58, CD104 and CD140a.The CD marker profile on PLA cells was further confirmed using flowcytometry. Osteogenic and adipogenic differentiation did notsignificantly change the CD profile, but, as with control cells, subtledistinctions could be determined using flow cytometry. Differentiationalso resulted in changes to the associated ECM. Finally, both ADSCclonal populations expressed a similar complement of CD antigens to thatobserved in the heterogenous PLA population, suggesting that clonalisolation of a multi-lineage population from the PLA does not affect theexpression of cell surface proteins.

[0383] Characterization of a cell population can be performed throughidentification of unique proteins expressed on the cell surface. Severalgroups have subsequently characterized MSCs based on their expression ofcell-specific proteins (e.g. STRO-1, SH2, SH3, SH4) and “clusterdesignation” (CD) marker profiles (Bruder, S. P. et al., 1998, Clin.Orthop., S247-256; Conget, P. A. et al., 1999, J. Cell Physiol,181:67-73; Pittenger, M. F et al., 1999, Science, 284:143-147.) Thisstudy confirms that, like MSCs, a unique combination of cell surfaceproteins are expressed on PLA cells with the two populations showingsimilar expression profiles. Like MSCs, PLA cells expressed CD13, CD29,CD44, CD71, CD90, CD105/SH2 and SH3 as shown by a combination of IF andFC. In addition, PLA cells did not express CD14, CD16, CD31, CD34, CD45,CD56, and CD62E on the cell surface. The similarity in CD profiles toMSCs lends support to the theory that PLA cells are a stem cellpopulation. However, the degree of similarity may indicate that PLAcells are simply an MSC population located within or contaminating theadipose compartment. Lipoplasty results in the rupture of multiple bloodvessels and while vasoconstrictors are used to minimize blood loss, theprocessed PLA pellet may be MSCs obtained from the peripheral bloodsupply (Zvaifler, N.J. et al., 2000, Arthritis Res., 2:477-488.)However, there appear to be a few subtle distinctions between PLA andMSC populations. In contrast to MSCs, no expression of CD58 could bedetected on PLA cells using IF, while expression was seen on MSCs (FIG.23). Furthermore, MSCs have also been reported to express CD104, CD106and CD140a (Bruder, S. P. et al., 1998, Clin. Orthop., S247-256; Conget,P. A. et al., 1999, J. Cell Physiol, 181:67-73; Pittenger, M. F et al.,1999, Science, 284:143-147.) No expression of these CD antigens weredetected on PLA cells using IF or FC (FIGS. 23 and 24). Thesedifferences may indicate that the PLA population represents a distinctpopulation of stem cells. However, the possibility that PLA cells are aclonal variation of MSCs cannot be completely ruled out.

[0384] Multi-lineage differentiation by PLA cells may result from thecommitment of multiple lineage-specific precursors rather than thepresence of a pluripotent stem cell population within adipose tissue.Therefore, multi-lineage differentiation by clonal isolates derived fromsingle PLA cells is critical to the classification of PLA cells as asource of stem cells. In support of this, ADSC isolated exhibitedmulti-lineage capacity in vitro staining positively using the histologicassays alkaline phosphatase (osteogenesis), Oil Red O (adipogenesis) andAlcian Blue (chondrogenesis). Several lineage combinations wereobserved, including tri-lineage (osteogenic, adipogenic andchondrogenic), dual-lineage (osteogenic/adipogenic,osteogenic/chondrogenic) and single lineage (adipogenic only). Isolationand expansion of ADSCs did not alter the CD expression profile and nodifference in CD expression could be detected between any tri-lineageand dual-lineage ADSC population. Therefore, the presence ofmulti-lineage ADSC isolates and their identical CD marker profile toheterogenous PLA cells further supports the theory that the adiposecompartment is a source of multi-potential stem cells.

[0385] Differentiation of mesenchymal precursors and stem cells may leadto changes in the expression of several cell surface and intracellularproteins as these cells acquire a new fate and function. To assess this,undifferentiated PLA cells and cells induced toward the osteogenic andadipogenic lineages were examined by IF and FC for any changes in CDmarker profile. Osteogenic differentiation did not significantly alterthe CD profiles of PLA cells (FIG. 2 and Tables 16 and 17). Indirect IFconfirmed the expression of CD44 and CD90 and did not detect expressionof CD34 and CD45 in both osteogenic PLA and MSC cultures. In addition,both osteogenic PLA and MSC cultures were positive for CD29, CD71, CD105and SH3 expression, whereas no expression of CD31 could be detected.However, further analysis of osteogenic PLA cultures by FC revealedsubtle changes to the CD profile. Specifically, osteo-induction resultedin a 1.8-fold and 3.8-fold increase in CD90 and CD44 expression levels,respectively. The increased expression of CD44, the hyaluronan receptor,is likely the result of increased matrix synthesis and cell-matrixinteraction by PLA cells upon osteogenesis. Recent work has alsoconfirmed the expression of Thy-1/CD90 on osteoblasts andosteoblast-like cells derived from mice, rats and human. Expression ofthis protein increased markedly during the earliest stages of maturation(proliferative phase) and decreased as the osteoblasts matured. Theincreased expression of CD90 upon osteogenic induction of PLA cells may,therefore, reflect the increased expression of this protein as theosteogenic PLA cells proliferate during the earliest phases ofdifferentiation. In addition to CD44 and CD90, a dramatic increase(6.2-fold) in the metalloprotease, CD13/aminopeptidaseN, was alsoobserved in osteogenic PLA cells. In addition to its expression oncommitted progenitors of granulocytes and monocytes [Kishimoto, 1997#1082], CD13 has also been identified on fibroblasts, bone marrowstromal cells and osteoclasts (Syrjala, M. et al., 1994, Br. J.Haematol., 88:679-684). Recent work has identified an increase in CD13mRNA levels upon cell-cell contact (Kehlen, A. et al., 2000, J. CellBiochem, 80:115-123; Reimann, D. et al., 1997, J. Immunol.,158:33425-3432.) The dramatic increase in CD13 on PLA cells maytherefore be due to the increased cell to cell contact within osteogenicPLA cultures. Additionally, increased expression of proteases, such asCD13, on stem cells may also participate in differentiation by degradingregulatory peptides and proliferation agents that may affect thedevelopment of these cells (Young, H. E. et al, 1998, Wound RepairRegen, 6:66-75; Young, H. E. et al., 1999, Proc. Soc. Exp. Biol. Med.,221:63-71.)

[0386] Interestingly, FC measured slight increases in CD34, CD56, CD49d,CD61 and CD105 expression upon osteogenic induction. With the exceptionof CD105, these markers were not expressed on undifferentiated PLA cellsand MSCs and their increase is likely the result of differentiation. A2.6-fold increase in CD34 expression was detected in osteo-induced PLAcultures. This increase was consistent with the appearance ofCD34-positive regions within 6steogenic PLA cultures as shown by IF(FIG. 23). A slight increase in CD34 was also observed upon IF analysisof osteogenic MSCs (FIG. 40). However, this increase appeared to be theresult of an overall enhanced expression level by all MSCs. Osteogenicinduction also resulted in a 1.8-fold increase in CD56 expression.Identified as neural cell adhesion molecule (NCAM), CD56 is expressed onhaematopoietic stem cells (Kishimoto, T. et al, 1997, Leucocyte TypingVI. White Cell Differentiation Antigens. (Hamden, Conn.: GarlandPublishing), mediating their adhesion with adjacent cells and thesurrounding matrix (Lanier, L. L. et al, 1991, J. Immunol,146:4421-4426; Lanier, L. L. et al., 1989, J. Exp. Med., 183:681-689.)Although its function has not been confirmed, CD56 may act in a similarmanner in non-haematopoietic cells. In support of this, osteoblastsexpress NCAM, using this adhesion molecule to mediate cell and matrixinteractions and leading to their differentiation (Lee, Y. A. et al.,1992, J. Bone Miner. Res., 7:1435:1466.) The osteogenic differentiationof PLA cells, therefore, may induce elevated levels of this CD proteinin order to regulate the increasing cell-cell and cell-matrixinteractions during differentiation. The same explanation can likely beapplied to the observed 3-fold increase in the α4 integrin, CD49d.Finally, a small increase in CD105 expression was measured on osteogenicPLA cells. Classified as a type III TGFβ3 receptor (Cheifetz, S. et al.,1992, J. Biol. Chem., 267:19027-19030), CD105 is expressed on a widevariety of cells, including endothelial cells, B-lineage precursors,MSCs and a subset of CD34+ cells isolated from peripheral blood(Rokhlin, O. W. et al., 1995, J. Immunol., 154:4456-4465; Majumdar, M.K. et al., 1998, J. Cell Physiol., 176:57-66; Barry, F. P. et al., 1999,Biochem. Biophys. Res. Commun., 265:134-139; Pierelli, L. et al., 2000,Br. J. Hematol., 108:610-620.) While little is know of this proteinduring bone development, expression of CD1 05 is thought to decrease asosteogenic precursors proceed toward terminal differentiation,disappearing on mature osteoblasts (Haynesworth, S. E. et al., 1992,Bone, 13:69-80.) Therefore, the expression of CD105 on osteogenic PLAand MSCs, as shown by IF, may indicate that these cells represent anearly stage in differentiation and have not reached their finaldifferentiation stage. Furthermore, the slight increase in CD105expression on PLA cells, as measured by FC, correlates to the increasein CD34 and may reflect the increase in a CD34⁺ subset within theosteogenic culture.

[0387] Adipogenic differentiation of PLA cells has been shown to resultin an expanded morphology, together with the accumulation of multiplelipid-filled intracellular vacuoles (Zuk, P. et al., 2001, TissueEngineering, 7:209-226.) As a result, adipo-induced PLA cultures are aheterogenous mixture of lipid-filled cells (i.e. mature PLA adipocytes)and more immature fibroblastic cells. Consistent with this, FCcharacterization of adipogenic PLA cultures demonstrated a shift towarda population of larger, more granular cells. IF analysis confirmed theexpression of CD29, CD44, CD71, CD90 and CD105 on adipogenic PLAs andMSCs (FIG. 41). While equivalent levels of CD29, CD71 and CD105 werefound on both fibroblastic and lipid-filled cells, lower levels of CD44and CD90 were observed in the mature PLA adipocytes. Contrary to PLAcultures, no such restriction could be detected by IF in adipo-inducedMSCs. While expression of CD90 appeared to be decreased in lipid-filledPLA cells, virtually 100% of the PLA fibroblasts stained brightly forCD90 and a 1.8-fold increase in this protein was measured using FC, alevel comparable to that measured in osteogenic cultures (1.75-fold).Contrary to CD90, expression levels of the CD44-positive PLA fibroblastsappeared to vary, with cell staining ranging from intense to little orno CD44. In support of this, FC confirmed a 48% decrease in CD44expression in adipogenic PLA samples. A decrease was also measured forCD13/aminopeptidase N and the decrease of these two proteins is likely areflection of the remodeling of the ECM to one more consistent withadipogenic tissue.

[0388] Like osteogenic PLA cells, FC confirmed the absence of CD14,CD16, CD31, CD45, CD62E and CD106 in adipogenic PLA cultures whileexpression of CD34, CD49d and CD61 were slightly elevated in thesecells. While FC did not detect a significant increase in CD45 uponadipogenic induction, a small percentage of PLA cells positive for thisprotein was observed upon IF analysis. The increased expression of CD34on adipogenic PLA cells was not as large as that measured uponosteogenesis and IF analysis confirmed CD34 expression by all PLAmorphologies. However, expression was restricted to cells with afibroblastic morphology. Weak expression of both CD34 and CD45 were alsodetected upon IF analysis of adipogenic MSCs with expression observed inboth fibroblastic and lipid-filled cells.

[0389] Differentiation of mesenchymal precursors to theirlineage-committed cell types (i.e. osteoblasts, adipocytes) isaccompanied by synthesis and remodelling of an ECM. Variation of ECMcomposition and organization gives each tissue its specificcharacteristics and participates in the differentiation and growth ofthe constituent cell types. For example, bone matrix consists ofinorganic hydroxyapatite together with an organic fraction comprised ofproteoglycans and collagens, with collagen type 1 making up the majority(approx. 90% of the organic fraction). Cartilage matrix consists mainlyof collagens type 2 and 10 and multiple sulfated proteoglycans.Adipogenic ECMs are comprised of multiple collagen subtypes (1 through6), laminin and fibronectin. Together, these collagens are a part of theunique extracellular environment of each tissue and are crucial to thesurvival and function of the component cells. Based on this, theexpression of ECM collagens were examined in both control and inducedPLA cells and MSCs.

[0390] Non-induced PLA cells and MSCs expressed CNI, CNIV and CNV (FIG.43), in addition to CNIII. Both CNI and CNIII exhibited similar stainingpatterns in both cell populations and osteogenic induction did not alterthe intracellular distribution of these collagens. Furthermore, aqualitative increase in CNI was observed in several PLA and MSC samples.A large volume of work confirms the role of collagen type 1 inosteogenic differentiation. For example, CNI levels increase during theearly stages of rat calvarial osteoblast differentiation and inhibitionof this collagen totally blocks osteogenic differentiation (Stein, G. S.et al., 1990, Faseb J., 4:3111-3123; Lynch, et al., 1995, Exp. CellRes., 216:35-45.) Factors that are known to affect osteogenesis, such asdexamethasone, vitamin D and the parathyroid hormone, can directlyaffect levels of CNI. Furthermore, bone marrow stromal cells maintainedon CNI matrices differentiate into osteoblasts in vitro and induce boneformation in vivo, an effect that is not seen on CNII, CNIII or CNVmatrices. Therefore, the synthesis of CNI in pre-induced andosteo-induced PLA cultures in consistent with the role of this collagenin osteogenesis. Moreover, the similarities in CNI expression observedin both osteo-induced PLA cells and MSCs suggests that similarmechanisms may function in the osteogenic differentiation of these celltypes.

[0391] In addition to CNI, expression of CNIV and CNV were also observedin both control PLA and MSC cultures, distributed in a fibrillar patternconsistent their secretion into the extracellular environment. Thepresence of these collagens is a vital component of the osteogenic ECMas expression of these collagens is observed in whole bone marrowstroma, the osteoblasts of newly forming bone and in STRO-1-positivecolony derived stromal cell lines. In contrast to induced MSC cultures,osteogenic induction of PLA cells appeared to significantly decreaseCNIV synthesis and completely inhibited CNV expression.

[0392] Adipogenic differentiation resulted in additional distinctionsbetween PLA and MSC populations. Like osteogenic cultures, adipogenicinduction of PLA cells resulted in an inhibition of CNV expression.Moreover, adipogenesis also resulted in the inhibition of CNI. No suchinhibition was seen in adipo-induced MSCs. Rather, a reduced level ofCNIV synthesis was observed in adipogenic MSC populations with all threecollagen types (I, IV and V) exhibiting a fibrillar, extracellularexpression pattern. While weak cellular expression of CNI was alsoobserved in lipid-filled MSCs, the expression of CNIV and CNV appearedto remain extracellular. Like MSC samples, adipo-induced PLA cells alsoexpressed CNIV. However, CNIV expression in these cells remainedintracellular and appeared to be expressed exclusively in lipid-filledPLAs.

[0393] Like osteogenesis, several lines of evidence suggest that ECMcomponents, such as collagens, participate in adipogenesis. First,changes in the ECM lead to morphologic and cytoskeletal alterations thatare required for the expression of lipogenic enzymes (Kuri-Haruch, W. etal., 1984, Differentiation, 28; Spiegelman, B. M. et al., 1983, Cell,357-666.) Second, expression of CNI, CNIII and CNIV varies dramaticallyupon differentiation of 3T3-L1 cells (Weiner, F. R. et al., 1989,Biochem., 28:4094-4099.) Lastly, the ECM of developing adipose tissue isorganized during differentiation, an event thought to be mediated by theadipocytes themselves (Nakajima, I. et al., 1998, Differentiation,63:193-200.) Fibroblasts and adipocyte precursors with a fibroblasticmorphology synthesize and secrete type I and III collagens, in additionto small amounts of the basement membrane collagen, type IV (Goldberg,B., 1977, PNAS, 74:3322-3325; Alitano, K. et al., 1982, J. Cell Biol.,94:497-505; Cryer, A. et al., 1982, Eur. J. Clin., Invest., 12:235-238;Kuri-Harcuch, W. et al., 1984, Differentation, 28; Liau, G. et al.,1985, J. Biol., Chem., 260:531-536.) As these cells begin todifferentiate changes occur in cell morphology, cytoskeleton and thelevel and type of ECM secreted (Napolitano, L., 1963, J. Cell Biol.,18:663-679; Aratani, Y. et al., 1988, J. Biol. Chem., 263:16163-16169;Weiner, F. R. et al., 1989, Biochem., 28:4094-4099.) These changes, inturn, may be a requirement for their terminal differentiation intoadipocytes.

[0394] To study the synthesis and distribution of ECM components uponadipogenesis, several preadipocyte cell lines have been developed,including several 3T3 variants (Green, H. et al., 1974, Cell, 3:127-133)and a clonal preadipocyte cell line from Japanese cattle (BIP cells)(Aso, H. et al., 1995, Biochem. Biophys. Res. Commun., 213:369-374.)Adipose conversion of BIP cells results in production of an ECM similarto adipose tissue in which adipocytes are interconnected by a fibrillarnetwork of collagens I, II, IV, V and VI together with an intracellularexpression of CNIII (Nakajima, I. et al., 1998, Differentiation,63:193-200.) Like BIP cells, adipo-induction of both PLA cells and MSCsresulted in a similar intracellular distribution of CNIII. Furthermore,fibrils of CNI, CNIV and CNV were also associated with adipogenic MSCsand appeared to be organized randomly. The expression of similarcollagens and their random organization in adipogenic MSCs is consistentwith that observed upon differentiation of preadipocytes and suggeststhat comparable ECM synthesis and remodeling may occur upondifferentiation of these stem cells.

[0395] However, the adipogenic differentiation of PLA cells presentsseveral differences to several preadipocyte cell lines and MSCs. Likepreadipocyte cells, including BIP cells from cattle, and 3T3 cells frommice, pre-differentiated PLA cells synthesize CNI and CNV. However,these collagens are no longer observed upon differentiation. Thedisappearance of CNI and CNV in adipogenic PLA cultures may represent aspecific remodelling pathway unique to these cells. In support of this,changes in the pericellular environment that occur duringdifferentiation can change the intracellular environment and thesecretion of MMPs that degrade the surrounding ECM. Low levels of CNIVare also produced by preadipocytes and a dramatic increase is observedupon adipogenesis (Aratani, Y. et al., 1988, J. Biol. Chem.,263:16163-16169; Nakajima, I. et al., 1998, Differentiation,63:193-200.) While a qualitative increase in CNIV is observed inadipogenic PLA cultures, its fibrillar distribution is lost and thecollagen is restricted to lipid-filled PLA cells. The change in CNIVexpression pattern in comparison to preadipocytes and MSCs remainsunclear.

[0396] While EM observations of mature fat cells have identified aCNIV-rich basement membrane associated with several other fibrillarcollagens (Chase, W. H., 1959, J. Ultrastruc. Res., 2:283-287; Barnett,R. J., 1962, L. W. Kinsell, ed., (Springfield, Ill.: Charles C. Thomas);Angel., A. et al., 1970, B. Jeanrenaud and Hepp, D., et ed. (Thiene,Stuttgard: Academic Press), mature fat cells do not synthesizecollagens. Moreover, adipogenic precursors lose the capacity forcollagen synthesis in vitro during the post-confluent differentiationstage. However, collagen synthesis is critical for terminal adipocytedifferentiation and triacylglyerol accumulation indicating that thepredifferentiation expression of an ECM determines their ultimatephenotype. Therefore, the predifferentiation expression of CNI, CNIII,CNIV and CNV by PLA cells and MSCs may serve to initiate theirdifferentiation program. As differentiation proceeds and the appearanceof lipid-filled cells (i.e. mature adipocytes) increases, the synthesisof these collagens ceases, resulting in a collagenous ECM unique toadipose tissue. This is likely the case with the MSC population.However, the absence of CNI and CNV in PLA cultures may be the result ofa direct inhibition of synthesis or a dramatic remodeling of the ECM.The precise time of collagen inhibition upon PLA adipogenesis and/or thepossible existence of agents involved in collagen degradation remainsunknown.

What is claimed is:
 1. An isolated adipose-derived stem cell (ADSC). 2.The stem cell of claim 1, which can be cultured for at least 15 passageswithout differentiating.
 3. The stem cell of claim 1, that ismultipotent.
 4. The stem cell of claim 3, that differentiates intomesoderm, ectoderm or endoderm.
 5. An adipose-derived stem-cell enrichedfraction (ADSC-EF) of an adipose tissue sample from a subject, saidfraction substantially free of adipocytes.
 6. The stem cell of claim 1which is human.
 7. The stem cell of claim 1, which is geneticallymodified.
 8. A defined cell population comprising a plurality of thecell of claim
 1. 9. The defined cell population of claim 8 which ishomogenous.
 10. The defined cell population of claim 8 which isheterogeneous.
 11. The defined cell population of claim 8 which isclonal.
 12. A progeny cell of the stem cell of claim 4, committed todevelop into a mesodermal cell.
 13. A progeny cell of the stem cell ofclaim 4, committed to develop into an ectodermal cell.
 14. A progenycell of the stem cell of claim 4, committed to develop into anendodermal cell.
 15. Tissue comprised of the stem cell of claim 4, anddifferentiated mesodermal cells.
 16. Tissue comprised of the stem cellof claim 4, and differentiated ectodermal cells.
 17. Tissue comprised ofthe stem cell of claim 4, and differentiated into endodermal cells. 18.A method of inducing mesodermal tissue comprising culturing the stemcell of claim 4 in a mesoderm-inducing medium.
 19. A method of inducingectodermal tissue comprising culturing the stem cell of claim 4 in aectoderm-inducing medium.
 20. A method of inducing endodermal tissuecomprising culturing the stem cell of claim 4 in a endoderm-inducingmedium.
 21. A method of forming tissue in a subject comprisingintroducing the progeny cell of claim 12, 13 or 14 into a subject in asufficient amount to form mesodermal, ectodermal or endodermal tissue insaid subject.
 22. A method of regenerating or repairing tissue in asubject comprising introducing a stem cell of claim 1, 12, 13 or 14 intoa subject in a sufficient amount to regenerate or repair tissue.
 23. Amethod for obtaining an adipose-derived stem cell-enriched fraction(ADSC-EF) comprising treating a sample of adipose tissue from a subjectto remove adipocytes forming an adipose-derived stem-cell-enrichedfraction (ADSC-EF).
 24. The adipose-derived stem-cell enriched-fraction(ADSC-EF) obtained by the method of claim
 23. 25. The adipose-derivedstem cells (ADSCs) obtained by separating said cells from the ADSC-EF ofclaim
 24. 26. The stem cells of claim 25, wherein said stem cells aremultipotent.
 27. The stem cells of claim 26, wherein said stem celldifferentiate into mesoderm, ectoderm, or endoderm.
 28. Anadipose-derived lattice comprising adipose tissue extracelluar matrixsubstantially devoid of cells.
 29. The lattice of claim 28 which issubstantially anhydrous.
 30. The lattice of claim 28 which is hydrated.31. A composition comprising the cell of claim 1 and a biologicallycompatible lattice.
 32. A composition comprising the cell of claim 1 andthe lattice of claim 29 or
 30. 33. Progeny of the stem cell of claim 3.34. A method of delivering a transgene to an animal comprisingintroducing the stem cell of claim 1 containing a selected transgeneinto a subject, such that the transgene is expressed in the subject. 35.A method of inducing the differentiation of the cell of claim 1,comprising culturing the cell in a suitable medium effective to inducedifferentiation under suitable differentiation conditions.
 36. Themethod of claim 35 wherein said medium is a conditioned medium of aspecific cell type.
 37. A method of inducing the differentiation of thecell of claim 1, comprising co-culturing the cell with a cell of desiredlineage.
 38. A method of conditioning culture medium comprisingcontacting the medium with the cell of claim
 1. 39. The cultured mediumobtained by the method of claim
 38. 40. A kit for obtainingadipose-derived stem cells (ADSCs) from adipose tissues of a subjectcomprising means for separating the ADSCs from the adipose tissue. 41.The kit of claim 40, further comprising a device for isolating adiposetissue from a subject.
 42. The kit of claim 40, further comprising amedium for inducing differentiation of the adipose-derived stem cells.43. The kit of claim 40, further comprising a medium for culturing theADSCs.